Angiogenesis induction using organoid-tissue modules: A platform for modular vessel construction

Abstract

Angiogenesis is essential for successful tissue regeneration, particularly in clinical contexts such as ischemic injury, wound healing, and reconstructive therapies. However, the establishment of functional vasculature remains a major limitation in organoid-based systems. In this study, we developed vascularized organoid tissue modules (Angio-TMs) by incorporating human umbilical vein endothelial cells (HUVECs) into scaffold-free, self-organized constructs. Remarkably, the inclusion of HUVECs at 1% of the total cell population was sufficient to generate highly reproducible and structurally stable Angio-TMs, which exhibited clear endothelial differentiation and vascular functionality both in vitro and in vivo. Furthermore, inhibition of transforming growth factor (TGF)-β signaling in Angio-TMs led to a 2.5-fold increase in vessel length density, demonstrating a substantial enhancement in angiogenic potential. These findings highlight Angio-TMs as a robust and modular platform for engineering vascularized tissues and underscore their translational relevance in regenerative medicine and tissue transplantation.

Introduction

The establishment of a functional vascular network is a critical prerequisite for successful tissue regeneration, as oxygen and nutrient diffusion is limited to within 100–200 µm from the nearest capillary in avascular tissues.^1–3 This vascularization requirement is particularly evident in clinical conditions such as diabetic foot ulcers and critical limb ischemia, where impaired perfusion leads to tissue necrosis, chronic wounds, or limb loss if vascular regeneration fails.^4,5 Likewise, in reconstructive surgery or in chronic ischemic wounds such as those seen in diabetic foot ulcers, large-volume soft tissue defects require robust vascular support to prevent central necrosis and ensure tissue survival. ⁶

Accordingly, extensive efforts in tissue engineering have focused on developing biomaterial scaffolds, angiogenic growth factor delivery systems, and cell-based strategies to promote effective and timely vascularization within engineered constructs. These approaches aim to recreate the physiological microenvironment necessary for endothelial cell recruitment, migration, and network formation, which are essential for the survival and integration of large-scale tissue grafts.^7,8 Among these, the co-culture of mesenchymal stem cells (MSCs) with endothelial cells such as HUVECs has emerged as a widely studied strategy due to its synergistic vascular-forming capacity. ⁹

Among the various sources of MSCs, adipose-derived MSCs (ADMSCs) have gained particular attention for vascularization strategies due to their accessibility, abundant yield, and robust secretion of proangiogenic factors.^10,11 Beyond their inherent multipotency, MSCs exhibit perivascular characteristics and secrete a wide range of angiogenic and tissue-regenerative factors, including VEGF, HGF, bFGF, and angiopoietin-1, which promote endothelial cell survival, migration, and tubulogenesis.^12,13 ADMSCs can not only modulate the inflammatory microenvironment, improving graft tolerance and integration, but also act as pericyte-like stabilizers, supporting endothelial lumen formation and vessel maturation.^14,15 This ability to enhance the formation of vascular networks is particularly advantageous for tissue-engineered constructs and in vivo transplantation. ¹⁶

Organoids provide physiologically relevant in vitro models that mimic native tissue structure and function. However, the absence of intrinsic vasculature remains a major barrier, restricting their growth, viability, and translational potential. Overcoming this limitation by engineering vascularized, scaffold-free organoids represents a key objective in advancing organoid-based regenerative therapies.

In light of these challenges, we focused on leveraging the synergistic co-culture of hADMSCs and HUVECs—a strategy previously shown to promote vascularization—for the development of scaffold-free, pre-vascularized organoid modules. In our approach, we utilized human ADMSCs as the primary source to develop vascularized tissue constructs. These ADMSCs were incorporated with HUVECs during the early fabrication step, enabling pre-vascular network formation and preserving the inherent angiogenic and supportive functions of the adipose-derived cells.

Vascularization remains a major challenge in organoid research, particularly for therapeutic applications, developing vascularized organoids is now a key research area in regenerative medicine in the field. The integration of perfusable vasculature is essential not only for sustaining large-scale 3D organoids in vitro, but also for ensuring their functional viability following transplantation. Therefore, the advancement of vascularized organoid platforms is increasingly regarded as a key enabler for the successful clinical translation of organoid-based therapies.^2,17

However, the majority of studies still rely on exogenous scaffolds, biomaterials, or external angiogenic cues to support cellular organization and vessel formation.^1,3 These materials, while often effective in promoting initial structure, may trigger host immune responses, degrade into bio-incompatible byproducts, or impair long-term cell survival and integration following transplantation. Even though scaffold-free strategies—such as spheroids or 3D aggregates composed of hADMSCs and HUVECs—have demonstrated an enhanced capacity to form pre-vascular structures in vitro and, in some cases, develop into functional vasculature upon transplantation, they still face several critical limitations. As these constructs grow in size, passive diffusion alone becomes insufficient to maintain adequate oxygen and nutrient delivery to the core, often leading to central necrosis and reduced cell viability over time.^18–20 Moreover, long-term maintenance of large spheroids in vitro typically requires dynamic culture systems or perfusion-based bioreactors, which can be technically complex, resource-intensive, and difficult to translate into scalable or clinically feasible approaches.

Clinically applicable tissue engineering requires vascularization that is not only biologically functional but also consistent and amenable to scalable manufacturing. Key architectural features—such as vascular sprouting, branching, alignment, and endothelial lining—must be tightly controlled to ensure functional and perfusable vascular networks.^21,22 However, existing spheroid-based approaches often suffer from batch-to-batch variability in size, cellular organization, and vascular performance, which hinders standardization for therapeutic use.

To address this challenge, we developed an Angio-Organoid-Tissue Module (Angio Organoid-TM) using human hADMSCs and HUVECs, specifically designed for therapeutic applications. This system supports intrinsic oxygen and nutrient diffusion and enables robust and scalable vascularized tissue production, as previously validated in our recent Acta Biomaterialia study demonstrating its efficacy in supporting vascularized tissue formation.

In this platform, angiogenesis is initiated at the Microblock (MiB) stage by integrating HUVECs with hADMSCs, which enables early endothelial sprouting. These MiBs are subsequently assembled into scaffold-free, pre-angiogenic Organoid-TMs. Notably, during this process, TGF-β signaling is suppressed, which facilitates sprouting activation in the early phase of angiogenesis. Furthermore, the system is optimized not only for sprouting but also for enabling directional endothelial outgrowth from HUVEC-containing MiBs toward neighboring hADMSC-only MiBs, indicating guided angiogenic migration and early vascular integration, thereby establishing a highly effective platform for engineered angiogenesis.

We anticipate that our platform can address a range of clinical conditions associated with insufficient vascularization. Diabetic foot ulcers, a common complication of diabetes, and critical limb ischemia (CLI), a severe form of peripheral arterial disease, frequently progress to tissue necrosis or ulceration due to inadequate perfusion. Current therapeutic strategies, including pharmacological agents, stem cell injections, and scaffold-based approaches, aim to promote tissue healing and regeneration. However, these methods often fail to achieve long-term efficacy due to limited cell survival, poor engraftment, and the absence of pre-formed microvascular structures.^23,24

The Angio-TM platform presents a clinically relevant strategy that actively induces the formation of functional microvascular networks. Within these constructs, hADMSCs secrete key paracrine mediators, including vascular endothelial growth factor (VEGF) and hepatocyte growth factor (HGF), which potentiate the angiogenic activity of co-cultured HUVECs and promote the assembly of structured vascular architectures.^25,26 Notably, the vascular networks emerge via intrinsic self-organization in the absence of synthetic scaffolds, thereby enabling rapid anastomosis with host vasculatures upon transplantation. This scaffold-free configuration is expected to reduce the risk of immune-mediated rejection and facilitate efficient tissue integration and regeneration.^27,28

This strategy not only overcomes the limitations of conventional scaffold-based and spheroid systems but also establishes a robust and clinically relevant foundation for generating vascularized tissues for therapeutic applications. Furthermore, it offers valuable insights into the regulation of vascular morphogenesis and provides a practical framework for the development of translational strategies in regenerative medicine.

Materials and methods

Cell culture

We used human adipose-derived mesenchymal stem cells (hADMSCs), which have advantages of high growth rate and easy accessibility compared to other adult mesenchymal stem cells, as a cell source. ²⁹ hADMSCs were isolated from human fat tissue. And, all experimental procedures were approved by the Institutional Review Board of the School of Dentistry, Seoul National University (IRB No. S-D20170038). GFP-HUVECs (ANGIO-PROTEOMIE, Boston, USA) were used in experiments. hADMSCs were cultured in growth medium composed of Dulbecco`s modified eagle medium (DMEM; Hyclone, Logan, USA), 10% fetal bovine serum (FBS; Hyclone), 1% antibiotic-antimycotic (AA; Gibco; Thermo Fisher Scientific, Massachusetts, USA). GFP-HUVECs were cultured in EGM-2 medium (Lonza, Basel, Switzerland) composed of 5% FBS, 0.4% hydrocortisone, 4% human fibroblastic growth factor (hFGF)-B, 1% VEGF, 1% R3-insulin-like growth factor (IGF)-1, 1% ascorbic acid, 1% human epidermal growth factor (hEGF), and 1% gentamicin sulfate-amphotericin (GA-1000). Cells were then expanded under standard culture conditions in a humidified atmosphere with 5% CO₂ at 37℃. And Cells were replaced with medium every 3 days until they reached 80%—90% confluency. And CTS™ TrypLE™ Select Enzyme (Gibco) is used to isolate cells from the plate for subculture or experimentation.

Fabrication of 3D cellular Microblock (MiB) and Angio-Organoid-TM (Angio-TM)

To fabricate 3D cellular Microblock, called Angio-MiB, cells were harvested with CTS™ TrypLE™ Select Enzyme (Gibco) and collected by centrifugation at 400g for 3 min. Cells were washed three times with phosphate-buffered saline (PBS) and DMEM or EGM-2. And cells reconstituted to 9.0 × 10⁵ cells/mL and 6.0 × 10⁵ cells/mL. Next, each 2 mL of cell suspension was seeded into the wells on AggreWell™ 400 and 800 plates (STEMCELL Technologies, Vancouver, Canada) to achieve an average density of 3000 and 500 cells/MiB. In addition, hADMSC-only MiBs were cultured in ITS medium (DMEM supplemented with 40 μg/mL l-proline (Sigma-Aldrich Co.), 100 μg/mL sodium pyruvate (Sigma-Aldrich Co.), 50 μg/mL l-ascorbic acid 2-phosphate (Sigma-Aldrich Co.), 1X ITS (Gibco), and 1X AA (Gibco)).

To delineate the role of the presence of GFP-HUVECs with hADMSC Angio-MiB were also fabricated in a similar way. After washing, cells were mixed with GFP-HUVEC and hADMSC cells in the ratio of 5:95. And, these plates were incubated 3 days at 37℃ for Angio-MiBs formation. Fabricated hADMSC-only MiB and Angio-MiBs were harvested by firmly pipetting media up and down to dislodge cellular MiBs from AggreWell™ 400 and 800 plates. Harvested MiBs were maintained in a DMEM or EGM-2 media, respectively. To confirm the size of hADMSC-only MiB and Angio-MiBs, harvested MiBs were observed and imaged using Juli-stage Imaging System (NanoEntek, Seoul, Republic of Korea) under the bright and fluorescent field. Then, ImageJ (NIH, Bethesda, MD, USA) was used to quantify their size distribution. Also, to fabricated MiB fusion, mixed the harvested 3000 cell MiBs and 500 cell MiBs in a ratio of 1:5. And then, these plates were incubated for 3 days at 37℃ for organoid aggregation. Additionally, to validate the RNA-seq data-based results, hADMSC-only MiB were cultured in EGM medium and designated as EGMMiB. These were subsequently used for ELISA and proteomics analyses.

After that, to fabricate Angio-TM, mixed the harvested 3000 cell MiBs and 500 cell MiBs in the ratio of 1:12, resulting in a total of 1.125 × 10⁶ cells. And, these plates were incubated for 21 days at 37℃ for formation. And the dissociation for their analysis was a 0.1% collagenase type I + 0.1% pronase solution. And to identify the cell composition according to their culture period, they were dissociated using 0.1% collagenase type I + 0.1% pronase solution and analyzed using FACSLyric™ Flow Cytometer (BD bioscience, CA, USA).

Enzyme-linked immunosorbent assay (ELISA)

Briefly, each group was set up with hADMSCs in the single cell state before MiBs were made, followed by MiBs made with an equal number of cells, and finally an equal amount of hADMSC as a control. The prepared samples were subjected to protein extraction using M-PER buffer (Thermo Fisher Scientific) as lysis buffer. The extracted proteins were loaded in 96-well plates, three wells for each group, and analyzed based on the manufacturer’s protocols the method using Human VEGF Quantikine^® ELISA (R&D Systems, Minnesota, USA), Human HGF Quantikine^® ELISA (R&D Systems) and human Fibronectin Quantikine^® ELISA (R&D Systems). For each ELISA, the amount of hADMSC used was 0.5 × 10⁶ cells for Fibronectin, 1.5 × 10⁶ cells for VEGF and HGF. Plates were read at 562 nm, and each protein level was calculated using the standard curve generated with the standard sample.

RNA-seq analysis

Total RNA was isolated with PureLink™ RNA Mini kit (Invitrogen, Carlsbad, CA, USA) following the manufacturer’s instructions. RNA purity and integrity were assessed using a NanoDrop8000 spectrophotometer (Thermo Fisher Scientific) and a 2100 Bioanalyzer (Agilent Technologies, Santa Clara, CA, USA). The mRNA sequencing libraries were prepared with Truseq Stranded mRNA kit (Illumina, San Diego, CA, USA). Cluster generation occurred in the flow cell on the cBot automated cluster generation system (Illumina), followed by sequencing on Novaseq 6000 sequencing system (Illumina). The sequencing raw data underwent quality checks using FastQC (Babraham Bioinformatics, Babraham Institute, Cambridge, UK). Qualified reads were trimmed by Trimmomatic version 0.39. The reads were mapped to the reference genome by Tophat (v2.0.13). The aligned results were added to Cuffdiff (v2.2.0) to identify differentially expressed genes. For library normalization and dispersion estimation, geometric and pooled methods were applied. Data analysis and graphic visualization were performed utilizing ExDEGA v.5.0.0 (eBiogen, Seoul, Republic of Korea). The differential expression of each gene was evaluated by calculating the log2 value. Genes meeting the criteria of a false discovery rate <0.05 and a |fold change|>2 were selected as upregulated or downregulated genes.

Proteomics data analysis

Protein extracted to boiling for 10 mins at 100°C by added lysis buffer (25 mM ABC, 4% SDS) final concentration 2% ³⁰ and centrifugation at 18,000 rpm for 10 min. Protein concentration was determined by the bicinchoninic acid (BCA) method. Protein samples (20 μg) were separated by 12% SDS-PAGE and in-gel digestion was conducted.

Gel fraction and enzyme digestion war performed described previously ³¹ gels ware divided and sliced into seven fractions according to molecular weight. Sliced gels were washed and destained with a 30% methanol and distancing solution (10 mM ammonium bicarbonate and 50% acetonitrile). After drying, gels were reduction with 10 mM dithiothreitol and alkylation of cysteines with 55 mM iodoacetamide. After the gels were washed with distilled water, tryptic digestion was performed in 50 mM ammonium bicarbonate at 37°C for 12–16 h. Tryptic peptides was dried then extraction with an extraction solution (50 mM ammonium bicarbonate and 50% acetonitrile containing 5% trifluoroacetyl acid (TFA)) and kept at 4℃ prior to mass spectrometry. For LC–MS/MS analysis, samples were dissolved in 0.5% TFA.

Liquid chromatography with tandem mass spectrometry (LC–MS/MS) analysis was performed according to the methodology presented in a previous report. ³² Tryptic digested samples were dissolved with 0.5% trifluoroacetic acid prior to further analysis. A 5 µL dissolved sample were onto a 100 μm × 2 cm nanoviper trap column and 15 cm × 75 µm nanoviper analysis column (Thermo Fisher Scientific) at a flow rate of 300 nL/min and were eluted with a gradient of 5%—40% acetonitrile over 95 min. All MS and MS/MS spectra captured by the Q Exactive Plus mass spectrometer (Thermo Fisher Scientific) were acquired in data-dependent top 12 mode. Peptide identification was based on monoisotopic mass selection, precursor mass tolerance of ±5 Da, fragment mass tolerance of ±0.8 Da, two missed cleavage and fixed modification of carbamidomethyl cysteine. Acceptance of individual spectra based on Mascot ion threshold score of 0.05 that is calculated specifically for each database search. The Peptide validator was used to calculate the false discovery rates (FDR) at 1.0% for peptide and protein matches above the identity threshold.

Histochemistry

To confirm the morphologies of Angio-MiBs or Angio-TMs, after harvesting, they were fixed into 4% paraformaldehyde (PFA) at 4℃ overnight. Samples were gradually dehydrated in alcohol and embedded in paraffin using a Leica TP 1020 automatic tissue processor (Leica Microsystems, Buffalo Grove, Illinois). Then, samples were sectioned in 4 μm thickness. These sections were then deparaffinized with xylene, rehydrated with ethanol at a graded decreasing concentration of 100%—70%, and finally washed with distilled water. The slides were stained with hematoxylin (Sigma-Aldrich Co., St. Louis, MO, USA) and eosin (Sigma-Aldrich Co.), washed with distilled water (dH₂O), and pathological changes were captured as bright-field images using the DM6B Digital Upright Fluorescence Microscope (Leica Microsystems).

Immunohistochemistry and immunofluorescence staining

Immunohistochemistry (IHC) and immunofluorescence (IF) for the detection of CD31, α-SMA (alpha-smooth muscle actin), and vWF (von Willebrand factor) distribution using light microscopy were performed. For the detection of antibodies, the samples were fixed in 4% PFA at 4℃ overnight, embedded in paraffin, then sliced into 4 µm thick sections. After deparaffinization, the sections were blocked and treated with anti-CD31 (Abcam, Cambridge, UK), anti-α-SMA (Abcam) and anti-vWF (Dako, Brüsseler Straße, Denmark) antibody. Peroxidase-based detection (DAB), blocking and diluent solution were used with the VECTASTAIN Universal Quick HRP Kit (Peroxidase), R.T.U. (Ready-to-Use; Vector Laboratories, Newark, CA, USA). In addition, immunofluorescence was performed with Alexa Fluor™ 488 and Alexa Fluor™ 568 (Invitrogen) as secondary antibodies, and nuclei were counterstained with Hoechst 33342 (Invitrogen). Alexa Fluor™ 488 and Alexa Fluor™ 568 were each diluted 1:400, and Hoechst 33342 was diluted 1:500. Brightfield images (×400 and ×1000) and fluorescence images (×200 and ×400) were acquired using the DM6B Digital Upright Fluorescence Microscope (Leica Microsystems).

3D imaging

To identify the conformation of Angio-MiB or Angio-TM, the samples were fixed in 4% PFA at 4℃ overnight, washed in 0.1% triton X-100 in PBS (called PBST), then pretreated for overnight at room temperature with 6% bovine serum albumin (BSA, Thermo Fisher Scientific) + 0.2% triton X-100 (Thermo Fisher Scientific) + 0.01% sodium azide (Sigma-Aldrich Co.) in PBS (called PBS-blocking buffer). The samples were incubated with anti-CD31 and vWF antibodies diluted in PBS-blocking buffer for 2 days at room temperature. After incubation, the samples were washed with PBST. Subsequently, the antigen-antibody complexes were incubated with fluorescent secondary antibody (1:200 dilution) and Hoechst 33342 (1:300 dilution) in PBS-blocking buffer for 2 days at room temperature. Following this, the samples were washed again with PBST. Then, the samples were immersed in a solution of 25% urea (Sigma-Aldrich Co.) and 65% sucrose (Sigma-Aldrich Co.) in water, followed by 3D imaging and fluorescence imaging using the Leica TCS SP8 confocal microscope (Leica Microsystems).

Cell functional test

Cells were seeded in 6-well plates (SPL Life Sciences, Pocheon-si, Republic of Korea) and cultured overnight. The next day, the cells were washed twice with serum-free medium and incubated in 3 mL of serum-free medium for serum starvation at 37°C for 18 h. After starvation, the cells were washed with 3 mg/mL BSA diluted in Hank’s Balanced Salt Solution (HBSS, Hyclone; working assay/wash buffer).

1,1′-Dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate (DiI)-Ac-LDL (Invitrogen) was diluted to a final concentration of 10 μg/mL in serum-free medium, and 1 mL of this solution was added to each well. The cells were incubated for 4 h at 37°C, washed three times with working buffer, and then fixed with 4% PFA at room temperature for 30 min. After fixation, the cells were washed twice with working assay/wash buffer. Cells were mounted using a DAPI-containing mounting medium (Vector Laboratories). The fluorescence images (×200) were acquired using the DM6B Digital Upright Fluorescence Microscope (Leica Microsystems).

For quantitative analysis by FACSLyric™ Flow Cytometer, the cells were resuspended in a mixture of 100 μL 4% PFA and 100 μL FACS buffer (2% FBS in PBS) after fixation with 4% PFA, prior to measurement.

All procedures were performed in accordance with the manufacturer’s instructions. And, to identified whether the GFP-HUVECs that constructed the Angio-MiBs retained their functionality, the samples were dissociated using a 0.1% collagenase type I + 0.1% pronase solution. Subsequently, DiI-ac-LDL was labeled as described above.

To determine whether the vascular structures formed within the Angio-MiBs originated from hADMSCs or GFP-HUVECs, we performed a FACS assay using GFP and CD31 markers. For the assay, Angio-MiBs were dissociated using the same method described above. The dissociated cells were blocked with 1 μL of IgG in 100 μL of FACS buffer. Subsequently, the cells were incubated with PE-conjugated CD31 antibody at 4°C for 1 h and 30 min. After incubation, the cells were washed with FACS buffer and analyzed using a FACSLyric™ Flow Cytometer (BD Biosciences).

Animal experiments

All animal procedures were approved by the Institutional Review Board of Seoul National University (SNU-200915-11-2) and were conducted in accordance with the Seoul National University Guidelines for the Care and Use of Laboratory Animals. A total of 25 NOD.Cg-Prkdcscid Il2rgtm1Wjl/SzJ (NSG) male mice (10 weeks) were randomly and evenly divided into five groups for each time point. The groups were classified based on the type of sample injected into the mice as follows: the control (Con) group received only Matrigel (Corning, New York, USA); the MiB group received hADMSC-only MiBs; the Angio-MiB group received Angio-MiBs; the Organoid-TM group received hADMSC-only Organoid-TMs; and the Angio-TM group received Angio-TMs.

Animals were anesthetized with isoflurane using an animal anesthesia system. After shaving the lateral abdominal region along the midline, the samples were subcutaneously injected into two sites on either side of the abdominal midline. The injection sites were monitored for 21 days. At each designated time point, the implanted sites were harvested and the collected samples were divided for downstream analyses: one portion was used for hemoglobin assay, while the other was fixed in 4% PFA at 4°C overnight for histological and immunostaining analyses. For immunostaining, sections were stained with antibodies against Ku80 (Cell Signaling Technology, Massachusetts, USA), GFP (Abcam), and CD31 (Abcam).

To assess the extent of blood vessel formation, hemoglobin content was quantified using the Hemoglobin Assay Kit (MAK115, Sigma-Aldrich Co.) according to the manufacturer’s instructions. Tissue samples were homogenized in PBS and centrifuged at 10,000×g for 15 min at 4°C to collect the supernatants, which were subsequently used for hemoglobin quantification. The assay employs an improved Triton™/NaOH-based colorimetric method in which hemoglobin is oxidized to a chromogenic product detectable by absorbance at 400 nm.

Inhibitors treatment

To evaluate the effect of TGF-β on angiogenesis in Angio-TMs, vascular formation was first confirmed within Angio-TMs after 3 days of Angio-MiB culture followed by 3 days of Angio-TM maturation. Subsequently, the TGF-β receptor I/II dual inhibitor LY2109761 (Selleck Chemicals, Texas, USA) was administered as a 7-day treatment. The experimental groups included Angio-TMs cultured in EGM media alone (Angio-TM (−)), a vehicle control group cultured in EGM media containing DMSO only, and a treatment group cultured in EGM media supplemented with LY2109761 (Angio-TM (+)). No significant difference was observed between the DMSO-only and Angio-TM (−) groups (data not shown). LY2109761 was dissolved with DMSO to prepare 5 mM stock solution. EGM medium was used to dilute, and the final concentration of 1 μM was employed.

To quantify the endothelial networks, the number of nodes (Nb nodes), number of junctions (Nb junctions), total master segment length (Tot. master segment length), total length (Tot. lengths), and total branching length (Tot. branching length) were measured for each of the nine sample using the Angiogenesis Analyzer for ImageJ software (https://imagej.nih.gov/ij/). ³³ Vessel length density (µm/µm²) was calculated by dividing the total vessel length by the analyzed area from the measured results.^34,35

Quantitative real-time polymerase chain reaction (qRT-PCR)

To verify the efficacy of the TGF-β receptor I/II dual inhibitor, real-time PCR was used to quantitatively detect the expression of EGFL7 (Epidermal growth factor-like domain multiple 7) and ITGB3 (Integrin subunit beta 3) in Angio-TMs. Total RNA was extracted using a TRIzol Reagent (Invitrogen), the concentration of which was determined from the optical absorbance at 260 nm. cDNA was synthesized on PrimeScript™ RT Master Mix (Takara, Kusatsu, Shiga, Japan). Real-time PCR was performed using a QuantStudio 3 Real-Time PCR System (Applied Biosystems, Massachusetts, USA) with a qPCRBIO SyGreen MiX Hi-ROX (PCR Biosystems, London, UK) used in each reaction. Gene expressions of EGFL7 and ITGB3 was evaluated by real-time PCR.

The sequences of the sense and anti-sense primers for EGFL7 (Forward 5′-TGC TGA TGT GGC TTC TGG TGT TG-3′ Reverse 5′-GGT GGT GAG GAA GGG CTG GTA C-3′), ITGB3 (Forward 5′-CGA AAA TAC CTG CAA CCG TTA CT-3′ Reverse 5′-TTG CCA GTG TCC TTA AGC TCT TT-3′) and GAPDH (Forward 5′-GGA GCG AGA TCC CTC CAA AAT-3′ Reverse 5′-GGC TGT CAT ACT TCT CAT GG-3′) were designed. All primers were designed using Primer-BLAST based on the GRCh38/hg38 reference genome provided by the UCSC Genome Browser. The mRNAs were normalized to the housekeeping gene GAPDH, and relative mRNA expression levels among groups were calculated using the ΔΔCT method. ³⁶

TUNEL assay

To demonstrate that Angio-TM is more favorable for cell survival than conventional cell aggregates under identical conditions, we generated spherical aggregates (hereafter referred to as “Aggregate”) using the same total cell number and ratio but with different morphologies. Aggregates were composed of 99% hADMSCs and 1% GFP-HUVECs, totaling 1.125 × 10⁶ cells, and were seeded into V-bottom 96-well plates (Thermo Fisher Scientific), followed by centrifugation at 500×g for 5 min to induce spheroid formation. These aggregates were then cultured under the same conditions as the Angio-TM for 6 days. These samples were fixed in 4% PFA at 4℃ overnight, embedded in paraffin, and sectioned to a thickness of 4 μm. The deparaffinized sections underwent apoptotic cell detection using ApopTag^® Peroxidase In Situ Apoptosis Detection Kit (EMD Millipore, Temecula, CA, USA) according to manufacturer’s instructions. Briefly, the sections were treated with proteinase K, followed by quenching endogenous peroxidase activity with 3% H₂O₂. Equilibrated sections were then incubated with TdT enzyme and digoxigenin-nucleotides. After treatment with peroxidase-conjugated anti-digoxigenin antibody, color development was achieved using diaminobenzidine peroxidase substrate. Methyl green was used for counterstaining. Following mounting, images were randomly captured using the DM6B Digital Upright Fluorescence Microscope (Leica Microsystems) at a magnification of 400×. For the comparison, samples were prepared in triplicate for each group. TUNEL-stained slides were imaged by a second investigator blinded to group allocation, who randomly captured five fields per sample. The acquired images were analyzed in a blinded manner using the cell counter function in ImageJ to quantify TUNEL-positive cells. Statistical analysis was performed using Student’s t-test.

Statistical analysis

Student’s t-test and One-way ANOVA using SPSS (version 10.10; IBM Corp., Armonk, NY, USA) was used to identify significant differences between the control and experimental groups. All values are reported as the mean ± standard deviation (SD), and a p-value of <0.05 was considered significant.

Results

Microblock (MiB) formation from hADMSCs and HUVECs induces early angiogenesis and enables Angio-Organoid-TM fabrication

To induce angiogenesis using the Organoid-Tissue Module (Organoid-TM), we employed our previously developed TRTP (Tissue-Reforming Technology Platform). ³⁷ We previously named microblocks made only with hADMSCs as MiBs, and we now designate those that induce angiogenesis through co-culture with GFP-HUVECs as Angio-MiBs. GFP-HUVECs were co-cultured with hADMSCs to generate Angio-MiBs containing endothelial cells, which were then mixed with MiBs composed solely of hADMSCs. This strategy allows the formation of angiogenesis while maintaining the overall architecture of the Organoid-TM (Figure 1(a)). This vascularized construction is hereafter referred to as the Angio-Organoid-TM (Angio-TM), representing a distinct therapeutic platform for inducing and sustaining angiogenesis within organoid-based tissue models. Previously, we observed that the Organoid-TMs exhibited distinct transcripts expression compared to hADMSC, revealing several intriguing differences in gene expression. Notably, the expression levels of angiogenesis-related genes—including ANGPTL4, VEGFA, ANG, and ITGB3—were significantly upregulated in the Organoid-TMs or MiB compared to monolayer-cultured hADMSCs. In particular, ANGPTL4 showed a 29.4-fold increase, followed by ANG (17.5-fold), ITGB3 (5.4-fold), and VEGFA (9.5-fold; Figure 1(b)). In this RNA-seq analysis, FN expression remained unchanged, whereas VEGF expression increased by 3.4-fold in MiBs and 17.5-fold in Organoid-TMs compared to single cells. Similarly, HGF expression rose by 6.6-fold and 22.4-fold, respectively. Based on these results, ELISA for fibronectin (FN), HGF, and VEGF were performed prior to Angio-TM fabrication to assess the proangiogenic profile of the MiB stage. We extracted proteins directly from the MiBs for the experiment. The elevated levels of these angiogenesis-related proteins further support the role of MiBs as a proangiogenic microenvironment that facilitates early vascular development within the Angio-TM system (Figure 1(c)).

Figure 1.

Fabrication of Angio-TM using hADMSCs and GFP-HUVECs: schematic and feasibility. (a) Schematic of Angio-Organoid-TM (Angio-TM) fabrication. Human adipose-derived mesenchymal stem cells (hADMSCs) and GFP-human umbilical vein endothelial cells (HUVECs) are seeded and aggregated over several days, forming Angio-MiBs and Angio-TMs. The figure was created with BioRender.com. (b) Volcano plots of MiB and TM compared to a single cell by RNA-seq analysis. (c) ELISA assay for VEGF, fibronectin, and HGF levels. Samples included hADMSCs, GFP-HUVECs, and three MiB types: ITS-based MiBs, EGM-based MiBs, and Angio-MiBs (co-cultured with GFP-HUVECs). Data represent mean ± SEM (n = 3). Statistical significance was determined by one-way ANOVA. Statistical significance: *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. (d) MiB fabrication using different cell types and media. hADMSCs and GFP-HUVECs were seeded and aggregated over 7 days to form MiBs in either ITS-based or EGM-based media. (e) Fluorescence images show the samples described in panel D over time. (f, g) Immunofluorescence (IF) staining detected endothelial differentiation through co-cultured GFP-HUVECs. Scale bar: 50 µm. (h) Three-dimensional image corresponding to panel G. Scale bars: 500 µm in panel E; 50 µm in panel F; 25 µm in panel G; and 50 µm in panel H.

To induce angiogenesis within Angio-MiBs, we co-cultured hADMSCs with GFP-HUVECs under standard culture conditions in well plates. As a control, MiBs composed solely of hADMSCs were cultured in EGM medium to assess whether the medium alone could induce angiogenic differentiation. After 7 days, the formation of endothelial alignment was observed exclusively in the Angio-MiBs (Figure 1(d) and (e)). During the initial phase of Angio-MiB formation, cells were uniformly dispersed in the wells; however, over time, the cells aggregated centrally within the wells, forming compact spheroids. Within these spheroids, GFP-HUVECs were localized to the periphery and exhibited a lining-like organization, suggestive of early vascular alignment. We analyzed Angio-MiBs containing 2% and 5% GFP-HUVECs using the Angiogenesis Analyzer plugin in ImageJ and found that the total vessel length and total branching length increased by approximately 160% and 211%, respectively, in the 5% group compared to the 2% group (Supplemental Figure 1B). This phenomenon was most prominently observed when 5% GFP-HUVECs were mixed with 95% hADMSCs (Figure 1(e)).

To determine the cellular origin of endothelial structures within Angio-MiBs, we performed CD31 immunofluorescence staining. MiBs composed solely of hADMSCs were cultured in endothelial growth medium (EGM) and included as controls. Across three independent experiments, hADMSC-only MiBs exhibited no detectable CD31 expression, regardless of media conditions (Figure 1(f)). Negative controls using secondary antibody alone confirmed staining specificity. These findings indicate that endothelial cells expressing CD31 arose exclusively from GFP-HUVECs, with no contribution from hADMSCs under these conditions. To further confirm the endothelial identity of the lining structures within Angio-MiBs, we performed immunofluorescence staining using human endothelial markers. Co-localization of CD31 and GFP indicated that the aligned endothelial architecture originated from incorporated GFP-HUVECs (Figure 1(g)). Three-dimensional imaging further revealed the spatial organization of these vascular structures (Figure 1(h), Supplemental Figure 1). To evaluate the generalizability of our approach, we also generated Angio-MiBs using fluorescently labeled human endothelial progenitor cells (hEPCs), which exhibited similar angiogenic features, supporting the robustness and versatility of the platform (Supplemental Figure 2).

To definitively determine the cellular origin of vascular structures, we conducted flow cytometry using GFP (FITC) and CD31 (PE), with isotype controls included. Analysis showed that 95.8% of CD31^⁺ cells were GFP^⁺, whereas only 0.1% were GFP^⁻ CD31^⁺ (Supplemental Figure 1D), confirming that endothelial structures were predominantly derived from GFP-HUVECs.

These results suggest that hADMSC-derived MiBs expressed key proangiogenic factors and, when co-cultured with GFP-HUVECs, effectively promoted angiogenesis—highlighting MiBs as a supportive niche for vascular formation.

Linking vessel-like structure from block to block

Since the Organoid-TM is a larger structure formed through the interaction between individual MiBs, we investigated whether angiogenesis could be initiated and extended when the construction was composed of both Angio-MiBs and hADMSC-only MiBs. Specifically, we examined whether new blood vessels would form and extend across the interface where Angio-MiBs were in contact with hADMSC-derived MiBs. We generated large MiBs composed solely of hADMSCs using 3000 cells per MiB, and smaller Angio-MiBs using a mixture of 95% hADMSCs and 5% GFP-HUVECs with 500 cells per MiB. To evaluate this, we co-cultured the two types of MiBs and monitored their interactions over time. In Figure 2(a), time-lapse microscopy of their mixing, the small Angio-MiBs moved the large MiB from 0 to 8 h. At 22 h, small Angio-MiBs were aggregated into large ones until 38 h. The small Angio-MiBs progressively appeared to aggregate around the large MiB and fused with it, resulting in their incorporation into the core of the larger MiB over time. At 46 h, GFP-positive endothelial structures within the Angio-MiBs were observed to extend toward the adjacent large MiB. Subsequently, GFP-expressing HUVECs exhibited an extending morphology, indicative of sprouting behavior toward the surrounding hADMSC-only MiBs (Figure 2(a)). These observations suggest that endothelial structures within Angio-MiBs are capable of directed extension toward neighboring hADMSC-only MiBs, supporting the concept of inter-MiB vascular integration within the Organoid-TM.

Figure 2.

Linking and extending endothelial alignment from one MiBs to others: (a) Time points of fusion step of MiBs. Angio-MiBs were generated by co-culturing hADMSCs and GFP-HUVECs for 14 days. Time-lapse images showing the fusion of a large hADMSC-only MiB (3,000 cells) with a smaller Angio-MiB (500 cells, 5% GFP-HUVECs). H&E staining of the fused MiBs shown in panel A. Fusion samples were stained with hematoxylin and eosin (H&E). Scale bar (Panelb): 50 µm. (c) Immunohistochemistry images of fusion MiBs. It was stained with α-SMA (brown) and CD31 (red). Scale bar (Panel c): each 50 and 25 µm, respectively. (d) Fluorescence images stained with CD31 (red) and GFP-HUVEC (green). (e) Three-dimensional reconstruction of the structures shown in panel d. (f) Fluorescence images stained with vWF (red) and GFP-HUVEC (green). (g) Three-dimensional reconstruction of panelf. In panels d andf, scale bars represent 50 µm; in panels e and g, 200 µm.

To determine whether the vascular structures extending from the Angio-MiBs into the hADMSC-only MiBs had undergone endothelial differentiation, we conducted immunohistochemistry. Immunostaining revealed co-localization of CD31 (red) and α-SMA (brown) within the same regions (Figure 2(c)), suggesting that the vascular structures formed in the Angio-MiBs were accompanied by the recruitment or differentiation of surrounding MSCs into pericyte-like cells.

Immunofluorescence analysis revealed that CD31 and vWF signals overlapped with GFP expression, indicating that the GFP-positive HUVECs contributed to endothelial differentiation. These signals extended from the Angio-MiBs toward the adjacent hADMSC-only MiBs and formed interconnected vascular structures (Figure 2(d) and (f)). Three-dimensional reconstructions further demonstrated the overall morphology and continuity of these vessel-like networks (Figure 2(e) and (g)).

Angio-TM-mediated vascular generation and elongation

To investigate whether sprouting vasculature could be induced within larger-scale constructs, we utilized the modular assembly of Organoid-TMs—previously shown to possess a cup-shaped geometry favorable for nutrient and oxygen diffusion—as a permissive microenvironment for vascular growth, characterized by sufficient porosity, favorable geometry, and cellular compatibility that collectively facilitate endothelial sprouting and network formation. Given our prior observation that vasculature formed in Angio-MiBs extended toward adjacent hADMSC-only MiBs, we hypothesized that similar interactions could promote interconnectivity and vascular integration at a across larger Organoid-TM structures. We therefore applied this principle to generate vascularized Organoid-TMs, hereafter referred to as Angio-Organoid-TMs (Angio-TMs). For Angio-TM fabrication, 3000 cells Angio-MiBs were prepared using a mixture of 5% GFP-HUVECs and 95% hADMSCs, while 500 cells MiBs consisted of only hADMSCs. As the proportion of GFP-HUVECs increased within the Organoid-TM constructs, we observed impaired formation of stable Organoid-TM structures. As shown in Figure 3(a), even at ~1.67% GFP-HUVECs, the formation of Angio-TMs was largely unsuccessful. Ultimately, a stable Angio-TM assembly was achieved when the GFP-HUVEC proportion was reduced to approximately 1%. According to Cho et al., ³⁷ the spatial occupancy of MiBs within the Organoid-TM structure is a critical factor influencing the formation of the characteristic architecture. Based on this, we observed that GFP-HUVECs, being smaller in size than hADMSCs, resulted in spheroids with reduced diameters when seeded at equivalent cell numbers, leading to an approximately 10% deficit in occupied volume. To address this, the total cell number was increased by 1.25-fold (1.125 × 10^⁶ cells), which effectively restored the structural morphology, with no significant improvement observed when further increased to 1.5-fold (1.35 × 10^⁶ cells). All three conditions maintained a constant GFP-HUVEC ratio of 1% (v/v; Figure 3(b)). To investigate whether the formation of a cup-shaped morphology is also advantageous for supporting internal vascularization in Angio-TMs, we conducted a TUNEL assay (Supplemental Figure 4). Angio-TMs and spherical aggregates (Aggregates) were prepared using identical total cell numbers and ratios. On day 6 of culture, TUNEL assay results showed a significantly lower number of TUNEL-positive cells in Angio-TMs compared to Aggregates, indicating improved cell survival. These findings suggest that the cup-shaped morphology is also essential for Angio-TM formation. Accordingly, we selected Angio-TMs composed of 1% GFP-HUVECs as the standard condition, as this group exhibited both robust vascularization and the successful formation of the cup-shaped structure that supports efficient diffusion of nutrients and oxygen.

Figure 3.

Optimal proportion of GFP-HUVECs for microvascular organization in Angio-TM: (a) Aggregation based on the proportion of HUVECs in Angio-TM. Scare bar (panel a): 1 mm. (b) Morphology of Angio-TMs based on their occupied area. Scare bar (panel b) : 1 mm. (c) Representative fluorescence images of Angio-TMs stained for CD31 (red) and Hoechst (blue), under different experimental conditions. Scare bar (panel c): 100 µm. (d) Three-dimensional reconstructions of Angio-TMs shown in panel C. Scare bar (panel d): 100 µm. (e) Fluorescence image of an Angio-TM constructed with 2% GFP-HUVECs. Scare bar (panel e) : 50 µm. (f) Corresponding 3D reconstruction of panel e. Scare bar (panel f): 200 µm.

To confirm the formation of vascular structures in the Angio-TMs produced under these optimized conditions, we performed immunofluorescence staining for GFP-labeled HUVECs and CD31, a human endothelial marker. Notably, when the pre-assembly culture period of Angio-MiBs was varied between 3 and 7 days, no significant difference in vascular integration within the Angio-TMs was observed (Figure 3(c)). Immunofluorescence analysis revealed co-localization of GFP and CD31 in both the 1% (Figure 3(c) and(d)) and 2% (Figure 3(e) and (f)) GFP-HUVEC groups, indicating the formation of interconnected and extended vessel-like structures.

To confirm that functionally mature vascular structures emerge by day 21, we performed CD31 IF staining. The results revealed that CD31-positive networks extensively spanned the interior of the Angio-TMs, indicating a mature and well-integrated vasculature at this time point. These CD31-positive structures extended directionally across the Angio-TMs, suggesting a continuous, aligned vascular elongation rather than typical sprouting angiogenesis.

Although Angio-TMs containing 2% GFP-HUVECs failed to form the desired structure, we examined whether vascular differentiation still occurred under this condition. Immunofluorescence analysis confirmed the formation of vascular structures (Figure 3(e) and(f)). In contrast, Angio-TMs with a maintained 1% GFP-HUVEC ratio exhibited both robust vascularization and successful formation of the characteristic cup-shaped morphology, favorable for nutrient and oxygen diffusion, as shown in Figure 3(c), (d), and Supplemental Figure 4. Therefore, we determined that cup-shape is important for Angio-TMs for long-term, larger-sized cultures, and the ratio of GFP-HUVECs in Angio-TMs was determined to be 1%. These results demonstrate that the TRTP-based organoid-tissue module platform can be effectively applied to support vascularization. The optimal condition for generating Angio-TMs was identified as containing 1% GFP-HUVECs, under which robust vascular formation characterized by both sprouting and directional extension was observed by day 21.

Induction of functional vascular cells from HUVECs using Angio-TMs

To confirm that GFP-HUVECs retained their endothelial properties after forming vascularized structures within Angio-MiBs or Angio-TMs, we performed functional validation using DiI-labeled acetylated low-density lipoprotein (DiI-Ac-LDL) uptake. DiI-Ac-LDL is internalized via scavenger receptors, which are highly expressed on endothelial cells, resulting in strong red fluorescence within the cytoplasm. Because this uptake reflects scavenger receptor–mediated endocytosis, it serves as a reliable indicator of endothelial functionality.

GFP-HUVECs were incubated with DiI-Ac-LDL and analyzed by both fluorescence microscopy and flow cytometry (FACS). hADMSCs, which lack scavenger receptor activity, were used as a negative control. As expected, hADMSCs did not take up DiI-Ac-LDL. In contrast, DiI-Ac-LDL uptake by GFP-HUVECs confirmed their endothelial identity. Fluorescence microscopy revealed intense red cytoplasmic staining, often clustered around the perinuclear region, while nuclei were counterstained with DAPI (blue; Figure 4(a)).

Figure 4.

Functional studies for identification of endothelial cells: (a) DiI-ac-LDL assay for endothelial cells compared to hADMSC. DiI-ac-LDL was taken up by endothelial cells and detected in the cytoplasm of the cell. hADMSC was used as a negative control. Nuclei were counterstained with DAPI (Magnification x200). Scare bar (panel a): 50 µm. (b) Flow cytometry analysis of DiI-ac-LDL uptake. Dot plot confirm functional DiI-ac-LDL internalization by endothelial cells compared to hADMSCs (Magnification ×200). Scare bar (panel b): 100 µm. (c) DiI-ac-LDL uptake assay in Angio-MiBs, confirming the functional capacity of endothelial cells forming endothelial alignment within the 3D spheroids. (d, e) Quantification of endothelial cells in Angio-MiBs by fluorescence-activated cell sorting (FACS). Angio-MiBs are dissociated into single cells and stained for endothelial markers. The proportion of endothelial cells was quantified to confirm integration within the construct. Data represents mean ± SEM (n = 3). Statistical significance was determined by one-way ANOVA, **p < 0.01.

In Figure 4(a), hADMSCs exhibited fluorescence only for DAPI, confirming the absence of both GFP expression and DiI-ac-LDL uptake. In contrast, GFP-HUVECs showed positive signals for GFP, DiI-ac-LDL (PE), and DAPI, indicating endothelial identity and functional LDL uptake. To quantitatively evaluate this uptake, we performed FACS analysis using DiI-ac-LDL as a marker of endothelial function (Figure 4(b)). hADMSCs, which lack LDL uptake capability, did not shift from baseline fluorescence in the PE channel, and their fluorescence profile overlapped between unlabeled (grey) and DiI-labeled (red) samples. In contrast, GFP-HUVECs demonstrated a distinct PE-positive signal, clearly separated from the negative control population, confirming their selective LDL uptake. Furthermore, additional analyses including cellular morphology, proliferation rate, and tube formation assay supported the endothelial characteristics of GFP-HUVECs (Supplemental Figure 5). These results confirm that GFP-HUVECs retained their functional characteristics as endothelial cells. To determine whether GFP-HUVECs retained their endothelial properties within Angio-MiBs during angiogenesis, Angio-MiBs cultured for 3 days were dissociated into single cells and replated in 6-well plates. The re-cultured cells were treated with DiI-ac-LDL, and fluorescence microscopy confirmed that GFP-HUVECs continued to uptake LDL, displaying red cytoplasmic staining indicative of functional endothelial activity (Figure 4(c)).

To investigate whether vessel extension in Angio-TMs was driven by migration or proliferation of GFP-HUVECs, we performed time-course FACS analysis. Prior to analysis, Angio-TMs cultured for 6, 21, and 28 days were enzymatically dissociated into single cells. On day 6, the proportion of GFP-HUVECs remained at approximately 1%, consistent with the initial seeding ratio. However, by days 21 and 28, the GFP-HUVEC population increased to 1.6% and 4.2%, respectively (Figure 4(d) and (e)), indicating proliferation of HUVECs during vascular network formation. These findings suggest that proliferation of GFP-HUVECs contributes to the expansion of endothelial structures within the Angio-TMs.

Based on the results from Angio-MiBs and Angio-TMs, the observed functionality of HUVECs as endothelial cells, along with their gradual proliferation over time, suggests both the angiogenic potential and expansion capacity of the system.

Assessment of angiogenesis capability of Angio-MiBs and Angio-TM in an in vivo Matrigel plug assay

To evaluate whether the vessel-like structures formed within Angio-TMs could function as blood vessels in vivo, a Matrigel plug assay was conducted using NSG mice (Figure 5). NSG mice were selected due to their immunodeficient status, allowing successful xenograft engraftment and survival of hADMSCs and GFP-HUVECs by reducing immune rejection. Mice were subcutaneously injected with Matrigel containing PBS (control), or Matrigel mixed with MiBs or TMs suspended in EGM-2 medium (Figure 5(a)). After 21 days, the Matrigel plugs containing Angio-MiBs appeared faintly reddish, while those containing Angio-TMs displayed distinct red microvascular organization visible on gross examination (Figure 5(b)). Hemoglobin content, measured using an improved Triton^TM/NaOH-based colorimetric method at 400 nm, significantly increased in the Angio-MiB and Angio-TM groups compared to controls, with the highest levels observed in the Angio-TM group (Figure 5(c)). The presence of human cells within the Matrigel was confirmed in all experimental groups (except control) via immunostaining for the human-specific nuclear marker Ku80 (Figure 5(d)). To confirm that the red structures represented blood vessels formed by the implanted GFP-HUVECs, we performed immunofluorescence staining for GFP and the endothelial marker CD31. Dual-positive signals for GFP and CD31 were observed only in the Angio-MiB and Angio-TM groups, with markedly stronger and more extensive staining in the Angio-TM group (Figure 5(e)), indicating enhanced endothelial integration and vascular formation.

Figure 5.

Assessment of angiogenesis capability of Angio-MiB and Angio-TM in vivo Matrigel plug assay: (a) Representative image showing the injection of Angio-TMs mixed Matrigel into the subcutaneous abdominal region. Samples with Matrigel were injected subcutaneously into abdominal regions in mice. (b) The morphology of the Matrigel retrieved after 21 days. (c) Hemoglobin assay for samples injected into mice. Assessment of angiogenesis capability inside Matrigel plugs by measurement of hemoglobin content. Data represents mean ± SEM (n = 3). Statistical significance: **p < 0.01. (d) Detection of the human-specific nuclear marker Ku80 (red) by immunofluorescence (Magnification ×400). Scale bar (panel d): 100 µm. (e) Fluorescence images stained with CD31 (red) and GFP-HUVEC (green; Magnification ×400). Scale bar (panel e): 50 µm.

Collectively, these findings demonstrate that while both Angio-MiBs and Angio-TMs possess angiogenic activity in vivo, Angio-TMs exhibit significantly enhanced vascularization potential. Notably, constructs composed solely of hADMSCs did not result in detectable vessel formation, underscoring the necessity of endothelial cell incorporation for functional vascular development.

Expression of genes regulated by TGF-β in Angio-TM

TGF-β is known to modulate angiogenic responses. To investigate its role in Angio-TMs, we treated the constructs with LY2109761, a TGF-β receptor I/II dual inhibitor, for 7 days following fabrication (Figure 6). One day after treatment, GFP expression in Angio-TMs appeared more intense, sharply defined, and aligned compared to untreated controls. After 1 week, GFP signals became more widely distributed and extended throughout the Angio-TMs, indicating enhanced vascular outgrowth (Figure 6(a)). To evaluate whether TGF-β signaling inhibition enhances vascular network complexity and spatial expansion, we conducted quantitative analysis using the Angiogenesis Analyzer.^33,38,39 Compared to the untreated group (−), the LY2109761-treated group (+) showed a significant increase in the number of nodes and junctions, indicating greater interconnectivity within the vascular network (Figure 6(b)). The master segment length, which reflects network complexity, also increased slightly but significantly (Figure 6(b)). In addition, both total vessel length and total branch length were markedly higher in the LY2109761-treated group than in the control group (Figure 6(b)). Furthermore, vessel density, derived from the total vessel length and corresponding analyzed area quantified in Figure 6(b), exhibited an increase of up to 258% in the TGF-β inhibitor-treated group (+ group), corresponding to a ~2.5-fold enhancement relative to the untreated (–) group, as shown in Figure 6(c). These results demonstrate that TGF-β inhibition promotes the formation of a more highly interconnected vascular network within the Angio-TM.

Figure 6.

The suppression of TGF-β during angiogenesis in Angio-TM: (a) Image of changes in Angio-TM over 7 days of TGF-β receptor I/II dual inhibitor LY2109761 treatment (Magnification ×40). Scale bar (panel a): 500 and 100 µm. (b) Quantification of endothelial networks using the Angiogenesis Analyzer plugin in ImageJ, showing the number of nodes (Nb nodes), number of junctions (Nb junctions), total master segment length (Tot. master segment length), total length (Tot. length), and total branching length (Tot. branching length) compared between non-treated (–) and LY2109761-treated (+) groups. All lengths measured are in micrometers (µm). (c) The amount of angiogenesis was quantified vessel length density, expressed as micrometers of vessel length per square micrometer of area of effect (µm/µm²). Data represents mean ± SEM (n = 9). (d) Comparative analysis of EGFL7 and ITGB3 mRNA expression levels on Angio-TM treated TGF-β receptor I/II dual inhibitor LY2109761 compared to non-treated Angio-TM. Data represents mean ± SEM (n = 3). (e) Detection of the CD31 (brown) by IHC staining (Magnification ×50 and ×200). Scale bar: 100 µm. Statistical analysis was performed using Student’s t-test.

These findings indicate that suppression of TGF-β signaling enhances vascular extension within the Angio-TM system. The untreated group (–) served as the reference and was normalized to 1.0, with fold changes in the LY2109761-treated group (+) calculated relative to this baseline. Consistently, mRNA expression of the angiogenesis-related genes EGFL7 and ITGB3, which regulate endothelial sprouting and migration, was significantly upregulated upon TGF-β inhibition (Figure 6(d)).

To validate the extent of angiogenesis at the protein level, we performed immunohistochemical staining for CD31 following TGF-β inhibition. CD31^⁺ signal was markedly increased in LY2109761-treated Angio-TMs (+) group compared to (−) group (Figure 6(e)). These findings reinforce the role of TGF-β suppression in promoting angiogenesis within the Angio-TM.

Discussion

In tissue engineering, various strategies have been employed to create three-dimensional (3D) microenvironments that closely mimic in vivo conditions and preserve the regenerative properties of mesenchymal stem cells (MSCs). Unlike conventional two-dimensional cultures, 3D platforms more accurately reproduce spatial oxygen gradients, nutrient diffusion, and paracrine signaling,^40,41 making them promising tools for regenerative medicine and translational research. ⁴² However, one of the fundamental challenges in large-scale 3D constructs is central necrosis due to limited diffusion of oxygen and nutrients,^43,44 necessitating the formation of functional vasculature.

Angiogenesis—the formation of new blood vessels—is a physiological prerequisite for effective tissue regeneration and functional integration of grafts. Its role is particularly vital in ischemic diseases, wound healing, and organoid development,^7,45 and its dysregulation underlies multiple pathologies such as cancer and diabetic complications. Recent research on vascularized organoids, spheroids, 3D bioprinting, and dynamic culture systems has significantly advanced tissue engineering; however, key limitations remain. Vascularized spheroids and organoids frequently experience oxygen and nutrient deprivation, leading to central necrosis and high batch-to-batch variability. Moreover, they often fail to establish functional, perfusable vasculature after implantation.^46,47 While 3D bioprinting enables precise architectural control, it struggles to reproduce capillary-scale microvessels and sustain cell viability. ⁴⁸ Dynamic culture systems improve perfusion but still face challenges in mimicking physiological microenvironments. ⁴⁹ Despite recent progress, most existing vascularization models either rely on synthetic scaffolds with unknown long-term compatibility or show limited endothelial cell functionality due to poor structural support. ²

To address these limitations, we developed a scaffold-free vascularization strategy using the Organoid-Tissue Module (Organoid-TM) platform derived from self-organizing human adipose-derived MSCs (hADMSCs), originally proposed in Cho et al. ³⁷ By applying our previously established TRTP (Tissue-Reforming Technology Platform) to vascular tissue engineering, we generated Angio-MiBs by incorporating GFP-HUVECs into hADMSC-based MiBs. These Angio-MiBs exhibited elevated angiogenic protein secretions, including fibronectin (FN), HGF, and VEGF, and supported the formation of lining-like endothelial arrangements. These factors exhibit high expression from the Angio-MiB stage, prior to Angio-TM formation, and promote early vascular development by enhancing endothelial cell migration and inducing tip cell formation. ⁵⁰ Notably, the synergistic action of HGF and VEGF drives vascular network expansion and sprouting during Angio-TM formation.^51,52

RNA-seq analysis confirmed that Organoid-TMs formed from MiBs upregulated angiogenesis-associated genes such as ANG, ANGPTL4, VEGFA, and ITGB3. Notably, expression of ANGPTL4 and ITGB3 in RNA-seq was increased by 29.4-fold and 5.4-fold, respectively, when compared to MiBs alone, underscoring the architectural influence of Organoid-TMs on vascular activation. ANGPTL4 regulates endothelial cell metabolism, promotes vascular permeability, and enhances endothelial activation. It plays a crucial role in angiogenesis by augmenting sprouting angiogenesis, particularly through its interaction with VEGF signaling. ⁵³ ITGB3 functions as a key mediator by binding to the extracellular matrix and activating multiple signaling pathways that stimulate endothelial cell activation, migration, and survival, thereby strengthening sprouting angiogenesis.^54,55 Furthermore, both ANGPTL4 and integrin β3 (ITGB3) signaling pathways have been reported to converge on the PI3K/Akt axis, which is also a critical downstream effector of the Tie2 receptor pathway. ANGPTL4 can activate PI3K/Akt signaling through its interaction with integrins such as α5β1 and β3, promoting endothelial cell survival and vascular barrier integrity.^56,57 Moreover, ITGB3 has been shown to form a functional complex with Tie2 and modulate its downstream activation. ⁵⁸ Based on these findings, we speculate that ANGPTL4 might also influence Tie2-mediated angiogenic responses through shared downstream effectors. ⁵⁹ However, further mechanistic studies are warranted to validate this potential crosstalk.

To explore modular integration and vascular extension, we fabricated Angio-TMs by assembling Angio-MiBs with hADMSC-only MiBs. GFP-HUVECs within Angio-MiBs demonstrated directional extension into adjacent MiBs, forming interconnected CD31^⁺ and vWF^⁺ vessel-like networks. Immunohistochemistry confirmed α-SMA-positive perivascular stabilization, suggesting functional vessel maturation. Notably, varying HUVEC composition revealed that a 1% ratio yielded optimal vascular formation and TM integrity, while higher ratios disrupted module assembly. To confirm the cellular origin of the CD31^⁺ endothelial structures, we performed FACS analysis using GFP and CD31 markers. The results showed that the vast majority of CD31^⁺ cells were also GFP^⁺, indicating that the endothelial networks were predominantly derived from the incorporated GFP-HUVECs. Importantly, varying the HUVEC composition revealed that a 1% ratio yielded optimal vascular formation and TM integrity, whereas higher ratios disrupted module assembly. This observation aligns with previous studies indicating that excessively high proportions of endothelial cells (ECs) in spheroid or organoid cultures impair proper structural formation due to cell competition, spatial constraints, and altered paracrine signaling. ⁶⁰ Furthermore, other studies have demonstrated that when ECs dominate the cellular composition, increased homotypic adhesion and competition for limited space and resources hinder the spatial organization and integration of other essential cell types required for organoid maturation. ⁶¹ These conditions often lead to central necrosis or apoptosis within spheroids and disrupt the development of functional vascular networks. Therefore, maintaining an optimal EC ratio is critical for the generation of stable and functional 3D vascularized organoids or spheroids.

The inhibition of TGF-β signaling using LY2109761 enhanced endothelial sprouting and upregulated angiogenic markers such as EGFL7 and ITGB3, indicating that suppression of TGF-β signaling is required for promoting angiogenesis within the Angio-TM system (Figure 6(d)). Notably, quantitative analysis revealed that vessel length density increased by approximately 2.5-fold (a 258% increase) compared to the untreated control group, further supporting the pro-angiogenic effect of TGF-β inhibition. These findings underscore the pivotal role of TGF-β signaling as a regulatory axis in vascular morphogenesis. However, given its extensive crosstalk with other signaling networks and pleiotropic effects across various cell types, global inhibition of TGF-β may result in unintended consequences. Therefore, future studies should focus on identifying and targeting specific downstream effectors of the TGF-β pathway that directly govern angiogenic processes. This approach may allow for more precise and controlled modulation of vascularization in tissue-engineered systems.

This finding highlights the flexibility of the TRTP platform across tissue types, expanding its application beyond cartilage regeneration ³⁷ into therapeutic angiogenesis. Although direct quantitative comparison with other fabrication methods is challenging due to the limitations of image-based measurements in 3D organoid structures like Angio-TM, our scaffold-free Angio-TM system still offers distinct advantages in architectural reproducibility, self-assembly, and host integration potential compared to other MSC–endothelial strategies using fibrin beads, ⁶² nano-hydrogel scaffolds,^63,64 or spheroid cultures, ⁶⁵ our scaffold-free Angio-TM system enables superior architectural reproducibility, self-assembly, and host integration potential. Moreover, it addresses key translational limitations such as lack of standardization, batch variability, and inadequate in vivo persistence.

Our results demonstrated increased expression of key angiogenic factors—VEGFA, HGF, and fibronectin—within Organoid-TMs, which are likely to contribute to the formation and extension of vascular structures. Specifically, VEGFA acts as a pivotal regulator of sprouting angiogenesis by guiding the directional migration of endothelial tip cells and promoting filopodia extension ⁶⁶ . HGF regulates β1 integrin recycling via Arf6 GTPase signaling, thereby modulating endothelial cell adhesion and migration—processes essential for sprout formation and vascular network assembly.^52,67 In addition, fibronectin, a major component of the extracellular matrix (ECM), promotes endothelial cell adhesion, migration, and tube formation through its interaction with α5β1 integrin, supporting the structural stabilization of developing vasculature. ⁶⁸ While these findings suggest that the elevated secretion of these factors may underlie the vascular morphogenesis observed in our system, further mechanistic studies will be required to delineate their coordinated roles in sprouting, directional guidance, and vascular stabilization within Angio-TMs.

Collectively, our findings demonstrate that Angio-TMs represent a robust and scalable platform for the development of vascularized tissue constructs with potential translational applications in regenerative medicine. In the present study, subcutaneous implantation was employed as a preliminary in vivo model to confirm the feasibility of Angio-TM formation and maintenance under physiological conditions. While this model was not intended to assess therapeutic efficacy or vascular integration within a pathological microenvironment, it provided initial validation that the self-organized Angio-TMs can retain their structural integrity and pre-vascular features after transplantation. To better evaluate the therapeutic potential of Angio-TMs in clinically relevant settings, future studies will incorporate disease-specific models—such as diabetic wound healing or hindlimb ischemia—to determine their efficacy in promoting functional revascularization and tissue regeneration under ischemic condition.

Supplemental Material

sj-pptx-1-tej-10.1177_20417314251376104 – Supplemental material for Angiogenesis induction using organoid-tissue modules: A platform for modular vessel construction

Supplemental material, sj-pptx-1-tej-10.1177_20417314251376104 for Angiogenesis induction using organoid-tissue modules: A platform for modular vessel construction by Jin Ju Park, Eunjeong Seo, HyeRan Gwak, Jieun Lee, Hyun Ji Kim, Soyeon Jeong, Junhyung Kim, SangHyuk Lee and Jaejin Cho in Journal of Tissue Engineering