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. 2025 Sep 6;15(2):e02418. doi: 10.1002/adhm.202502418

Engineering Volumetric Tissue Analogs via Assembly of Endothelial Cell‐Covered Spheroidal Microtissues

Hyeongkwon Oh 1, Jae‐Hun Kim 2, Sunwoo Lee 1, Jin‐Hyung Shim 1,3, Won‐Soo Yun 1,3, Songwan Jin 1,3,✉
PMCID: PMC12805621  PMID: 40913564

Abstract

Tissue engineering holds a significant promise for the development of bioartificial organs applicable to transplantation. However, the size of engineered tissues remains limited, primarily due to the challenge of establishing microvascular networks within tissue constructs. In this study, engineered tissues are fabricated and embedded with functional microvascular networks by assembling endothelial cell‐covered spheroidal microtissues. Utilizing a preset extrusion bioprinting and microfluidic emulsification approach, spheroidal microtissues covered with endothelial cells are successfully fabricated, demonstrating high structural integrity compared with non‐structured spheroidal microtissues. Upon assembling endothelial cell‐covered microtissues, vascularized tissue structures are generated. Additionally, perfusion of the culture medium through the tissue structure created by microtissue assembly preserves the intermicrotissue spaces, which can be considered vascular channels, facilitating the development of highly vascularized tissues. By controlling the medium perfusion rate, volumetric functional tissues are produced that can be cultured for 7 days in vitro. This innovative technique for fabricating highly vascularized volumetric tissues holds potential as a foundational technology for producing artificial organs.

Keywords: core–shell, endothelial cell‐covered spheroid, microvascular network, perfusion culture, preset bioprinting, vascularized tissue, volumetric tissue

An engineering strategy to fabricate highly vascularized volumetric tissues with capillary networks is developed. Endothelial cell‐covered spheroidal microtissues with high structural integrity are efficiently produced using bioprinting technology. These spheroidal microtissues are then assembled and cultured under perfusion conditions, resulting in the successful fabrication of functional large‐volume tissues with high cell viability.

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1. Introduction

Tissue engineering has rapidly advanced over the past decades and emerged as a promising solution to the increasing shortage of organ donors for patients awaiting transplantation.[ 1 , 2 , 3 ] However, tissues produced using tissue engineering are often too small for clinical applications.[ 4 , 5 , 6 ] A major reason is the absence of vascularized structures, which are essential for cell survival by facilitating the transfer of nutrients and oxygen.[ 7 , 8 ] In the absence of vascular structures, the only mechanism by which essential substances are delivered to cells in tissue is by diffusion. In such cases, due to the diffusion limitation, hypoxia and nutrient deprivation can occur in large‐volume tissues, leading to cell necrosis.[ 9 , 10 , 11 ] Therefore, the development of vascularized tissue is crucial for fabricating larger and more functional tissues or organs.[ 12 , 13 , 14 , 15 ] Furthermore, when cells exist in tissue‐engineered constructs at a high density, the consumption of oxygen and nutrients increases. Consequently, cells far from the culture medium encounter greater challenges accessing the elements required for growth, underscoring the need to develop highly complex and intricate vascular networks.

The native liver produces blood components, such as albumin, and performs drug metabolism and detoxification.[ 16 ] A crucial factor in enabling these functions is the high cellular density, with cells comprising ≈80% of the total tissue volume.[ 17 ] In tissue engineering, replicating such a high cellular density is important. Lower cell density in tissue‐engineered constructs may result in functional differences compared to native tissues, rendering normal functional recovery impossible through organ transplantation.[ 18 , 19 ] These insufficient factors for interaction between cells in transplanted and native tissues may prevent stable engraftment and adequate cell signaling. Additionally, an excess of extracellular matrix components besides cells could lead to transplant failure, as the extracellular matrix cannot maintain its structure until cells proliferate sufficiently to enable tissue remodeling after transplantation.[ 20 , 21 ]

In tissue engineering, the conventional method for constructing tissue involves seeding cells that comprise the target tissue onto a scaffold, which serves as a 3D support for cell growth.[ 22 , 23 ] Subsequently, the scaffolds containing cells were cultured using a culture medium that included signals such as growth factors and cytokines to facilitate the completion of tissue formation. From a technical perspective, this approach is classified as top‐down tissue engineering,[ 23 , 24 ] and substantial research has been conducted to the extent that it can be applied to clinical applications. This approach is particularly effective for fabricating tissues such as skin[ 25 ] and cartilage,[ 26 ] which have low cell density,[ 17 ] simple internal architecture and cellular composition, or lack of vascularization,[ 27 ] allowing sufficient diffusion of essential nutrients within the tissue. Anderson et al.,[ 28 ] highlighted that significant efforts have been made to create functional tissue with high cell density and complex structures, such as liver tissue, by revitalizing decellularized extracellular matrix scaffolds. However, there are limitations to this approach. Most human tissues form highly organized and sophisticated structures, such as capillaries and hepatic lobules. Seeding or perfusing cells into scaffolds poses challenges for achieving precise cell placement and results in uneven cell density within the scaffold. Furthermore, it is difficult to incorporate highly vascularized structures capable of nutrient delivery within the scaffold. Consequently, cells located deep within the scaffold may not receive adequate nutrient transport, increasing the risk of necrosis.

In contrast, bottom‐up tissue engineering focuses on assembling small cellular units or aggregates, such as spheroids or organoids, into volumetric tissue structures.[ 29 ] A promising example of this approach is tissue fabrication via the assembly of cell spheroids. When spheroids come into contact with each other, they undergo fusion, leading to organized tissue structures that can be utilized for tissue construction. A representative example involves positioning cellular spheroids onto a needle array using 3D bioprinting to facilitate the fusion of adjacent spheroids for vascular tissue fabrication[ 30 ] or positioning spheroids at desired locations via aspiration to construct osteogenic tissue.[ 31 ] This fusion‐based tissue fabrication method enables the production of tissues with relatively high cell density by utilizing cell aggregates without scaffolds. Moreover, Skylar–Scott et al.,[ 32 ] successfully fabricated tissue structures with vascular channels that supply oxygen and nutrients by 3D printing sacrificial ink in a bath containing spheroids. While this approach enables the construction of large vessels, the challenge remains in densely arranging capillary‐like vascular networks.

In this study, we fabricated volumetric tissues with microvessel networks by assembling bioprinting‐based spheroidal microtissues. The key hypothesis of this study was that fusion between endothelial cell‐covered spheroidal microtissues would create intermicrotissue spaces that could function as a microvessel network. Therefore, we hypothesized that assembling endothelial cell‐covered spheroidal microtissues would enable the creation of tissue constructs with vascularized microchannels and that dynamic culture with medium perfusion could lead to the development of volumetric tissues. To validate our hypothesis, we employed a preset extrusion‐based bioprinting technique[ 33 , 34 ] to fabricate both core–shell structured spheroidal microtissues covered with endothelial cells and non‐structured spheroidal microtissues in which the constituent cells were uniformly mixed without a defined structure. These spheroidal microtissues were then cultured in vitro, and their structural integrity was evaluated. Furthermore, the differences in their structures were analyzed when the spheroidal microtissues were cultured adjacently within a confined environment. Subsequently, the bioprinted spheroidal microtissues were integrated at a moderate volume and subjected to dynamic culture conditions. Remarkably, although both groups underwent a tissue organizational process, the core–shell microtissue group maintained intermicrotissue spaces, which resulted in high cell viability. In contrast, the non‐structured microtissue group exhibited complete fusion of the microtissues without forming vascular channels, leading to cell necrosis. Building on these findings, we further expanded the tissue constructs by culturing larger assemblies of core–shell microtissues. Despite the large volume and high cell density of the tissue constructed through spheroidal microtissue assembly, cell viability and tissue functionality were successfully maintained during the 7‐day culture period. These findings highlight the potential for forming microvessel networks through bottom‐up tissue engineering using bioprinted core–shell microtissue, providing a promising pathway for developing clinically relevant and transplantable tissue constructs.

2. Results

2.1. Strategy for the Construction of Highly Vascularized Volumetric Tissues

Figure 1 shows the strategy for fabricating a highly vascularized volumetric tissue construct. First, bioinks were prepared by mixing EA.hy926 vascular endothelial cells with atelocollagen type I and HepG2/C3A parenchymal cells with atelocollagen and loaded into the shell and core compartments, respectively, of a precursor cartridge with core–shell architecture. The precursor cartridge containing the bioinks was transferred to a syringe and mounted onto a bioprinting system. Subsequently, pneumatic pressure was applied to the syringe to extrude both bioinks simultaneously through a nozzle. The atelocollagen used in bioinks as a hydrogel has a high viscosity, low extrusion velocity, and small nozzle diameter. Consequently, the extrusion process of the two bioinks operated under hydrodynamic conditions characterized by a low Reynolds number flow, preventing mixing and maintaining the core–shell structure of the precursor cartridge through to the nozzle exit. At the nozzle outlet, the aqueous‐phase bioinks met the oil phase (mineral oil), resulting in emulsification. This process formed spheroidal spheres while retaining the core–shell structure generated during extrusion, producing endothelial cell‐covered spheroids. We previously[ 34 ] reported that this process has a high throughput and fabricates over 45 spheroidal microtissues per minute, producing many core–shell spheroidal microtissues.

Figure 1.

Figure 1

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Schematic Representation of the Research Strategy. Volumetric vascularized tissue constructs were fabricated using core–shell structured spheroidal microtissue produced by combining preset extrusion bioprinting with a microfluidic emulsification system. The assembled microtissues were cultured in a 3D‐printed chamber under perfusion conditions to promote microvessel formation and ensure nutrient delivery throughout the tissue construct.

The bioprinted core–shell spheroidal microtissues were transferred into a 3D‐printed culture chamber, where voids naturally formed at the contact points between adjacent spheroidal microtissues. As these voids were uniformly distributed, they contributed to the development of a highly porous 3D structure. The culture medium was perfused using a pump to facilitate efficient oxygen and nutrient delivery within the tissue construct, allowing it to flow through the intermicrotissue spaces and support cell viability and tissue functionality. The spheroidal microtissues featured a core–shell structure with an endothelial cell‐covered surface layer. Consequently, the intermicrotissue voids functioned as microvessels, forming an extensive vascular network that enhanced tissue vascularization. During perfusion culture, spheroidal microtissue fusion occurred while maintaining the microvasculature channel, leading to the formation of a highly vascularized tissue construct. This approach enabled the generation of vascularized volumetric tissues, demonstrating the potential for engineered tissue development.

2.2. Structural Characteristics of Core–Shell and Non‐Structured Spheroidal microtissues

To evaluate the structural characteristics of individual spheroidal microtissues before vascularized tissue formation, both core–shell and non‐structured spheroidal microtissues were analyzed. Non‐structured spheroidal microtissues formed by randomly mixing HepG2/C3A and EA.hy926 cells without a defined organization were used as a comparison group. Figure 2A presents bright‐field images of bioprinted non‐structured and core–shell spheroidal microtissues at days 0, 4, and 14. The diameters of the microtissues were measured and are shown in Figure 2B. On day 0, immediately after bioprinting, both spheroidal microtissue types had similar diameters (non‐structured spheroidal microtissues had an average diameter of 410.1 µm, whereas core–shell spheroidal microtissues measured 412.8 µm). The diameters of both spheroidal microtissue types gradually decreased until day 4, reaching 288.5 µm for non‐structured spheroidal microtissues and 291.2 µm for core–shell spheroidal microtissues. However, a distinct divergence was observed after day 4. The non‐structured spheroidal microtissues increased in diameter over time, reaching an average of 352.2 µm by day 14. In contrast, the core–shell spheroidal microtissues maintained a diameter similar to that at day 4, measuring 280.1 µm. This trend is illustrated in Figure 2C, which depicts the diameter variation according to the spheroidal microtissue structure. By day 21, non‐structured spheroidal microtissues continued to increase in size, whereas core–shell spheroidal microtissues maintained a consistent diameter.

Figure 2.

Figure 2

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Structural Comparison of Core–Shell and Non‐Structured Spheroidal Microtissues Over Time. The comparison highlights the differences in morphology and stability. A) Bright‐field microscope images of non‐structured and core–shell spheroidal microtissues on days 0, 4, and 14. B) Quantification of diameter distributions. (n = 80). C) Diameter changes in non‐structured and core–shell spheroidal microtissues over time. (n = 30). D) Circularity measurements of non‐structured and core–shell spheroidal microtissues over time (n = 50). Data are presented as mean ± standard deviation (SD). E) Immunofluorescence staining of non‐structured and core–shell spheroidal microtissue surfaces and cross‐sections on days 1, 4, 7, 14, and 21.

Figure 2D further highlights the differences in structural stability, showing that core–shell spheroidal microtissues retained stable circularity throughout the culture period, whereas non‐structured spheroidal microtissues exhibited a progressive decrease in circularity. This suggests that core–shell spheroidal microtissues maintained their initial spherical morphology and structural integrity, whereas non‐structured spheroidal microtissues gradually lost their shape and became increasingly irregular.

The differences in diameter and circularity can be attributed to the structural organization of the spheroidal microtissues. Figure 2E presents the immunofluorescence staining of both spheroidal microtissue types over time to investigate these structural differences. The first and second rows display surface views of non‐structured and core–shell spheroidal microtissues, respectively, and the third and fourth rows show cross‐sectional views of their midsections acquired through confocal microscopy. Core–shell spheroidal microtissues exhibited a pre‐formed endothelial cell layer on their surface, which acted as a structural boundary, allowing them to maintain a spherical shape throughout the 21‐day culture period. In contrast, non‐structured spheroidal microtissues lacking this layer showed HepG2/C3A cells protruding through the surface by day 7. By day 14, these spheroidal microtissues had lost their initial spherical form and adopted an increasingly irregular morphology.

2.3. Comparison of Tissue Organization in Adjacent Core–Shell and Non‐Structured Spheroidal Microtissues

Building upon the structural characteristics observed in individual core–shell and non‐structured spheroidal microtissues, we next examined how these spheroidal microtissues behaved when cultured adjacently within a confined environment. Spheroidal microtissues were initially cultured individually until day 4, when their diameters stabilized. They were then positioned adjacent to each other on a Petri dish coated with poly (2‐hydroxyethyl methacrylate) (poly‐HEMA) to facilitate spheroidal microtissue aggregation. Figure 3 presents the immunofluorescence staining of spheroidal microtissue aggregates on days 1 and 14.

Figure 3.

Figure 3

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Immunofluorescence Staining of Spheroidal Microtissue Aggregates Showing the Structural Comparison of Core–Shell and Non‐Structured Spheroids on Days 1 and 14. Differences in morphology, stability, and cellular organization are observed. The first and second rows display surface views of aggregates formed from non‐structured and core–shell spheroidal microtissues, respectively. The third and fourth rows provide cross‐sectional views of the midsections, imaged using confocal microscopy. The second and fourth columns show magnified images of the regions outlined by white boxes in the first and third column images. The white arrows indicate the spatial separation between HepG2/C3A and EA.hy926 cells maintained in the core–shell spheroidal microtissue group.

On day 1, both core–shell and non‐structured spheroidal microtissues exhibited fusion and early tissue organization, with no significant differences observed in their surface views. However, cross‐sectional images revealed distinct structural differences between the two groups. In the non‐structured spheroidal microtissue group, HepG2/C3A and EA.hy926 cells were homogeneously distributed throughout the tissue. In contrast, a clear separation was maintained in the core–shell spheroidal microtissue group, with EA.hy926 cells forming an outer layer surrounding the HeG2/C3A cells, preserving their distinct spatial organization. By day 14, pronounced differences in tissue morphology emerged between the two groups. In the non‐structured spheroidal microtissue group, external images revealed irregular protrusions of HepG2/C3A cells extending outward, leading to the loss of the initial smooth, spherical surface. Internally, HepG2/C3A and EA.hy926 cells were irregularly distributed throughout the tissue. In contrast, the core–shell spheroidal microtissue group exhibited a smoother external surface. However, it remained intact, effectively preserving the spatial separation between HepG2/C3A and EA.hy926 cells without significant intermixing. These findings align with the structural differences observed in Figure 2, demonstrating that the preserved structural integrity of core–shell spheroidal microtissues contributed to the formation of more organized and stable tissue constructs, whereas non‐structured spheroidal microtissues exhibited irregular and disorganized growth over time.

2.4. Enhanced Structural Integrity and Viability of Core–Shell Spheroidal Microtissues Under Perfusion Culture

Previous results demonstrated that core–shell spheroidal microtissues exhibit superior structural stability compared to non‐structured spheroidal microtissues under both individual (Figure 2) and static culture conditions (Figure 3); however, it remains unclear whether these differences play a key role in the development of vascularized volumetric tissue. To address this, we designed and built a custom medium perfusion system to evaluate the structural differences between spheroidal microtissue‐based tissue constructs under perfusion culture conditions.

To implement perfusion culture, we developed a system that continuously supplies nutrients and oxygen to the tissue constructs. Figure 4A shows a schematic representation of the medium perfusion system designed for spheroidal microtissue‐based tissue constructs, in which spheroidal microtissues aggregate and circulate. Bioprinted spheroidal microtissues were placed inside a 3D‐printed culture chamber made of polycaprolactone (PCL), positioned at the center of a Petri dish. A peristaltic pump was used to circulate the culture medium. The unique microstructure of the culture chamber prevents spheroidal microtissues from escaping while maintaining a stable culture medium level within the chamber. Figure 4B presents a photograph of the actual 3D‐printed culture chamber and a scanning electron microscope (SEM) image highlighting its structural details. The chamber had an octagonal cross‐section with a side length of 1.66 mm and eight outlets at the bottom, each measuring 400 µm in width and 200 µm in height. This design prevented spheroidal microtissues from escaping the chamber while facilitating their accumulation inside. Owing to the small channel size of the 3D‐printed chamber and microporous structure formed by spheroid assembly, the culture medium was maintained at a constant level without complete leakage. When a sufficient hydrostatic pressure threshold is reached, culture medium flow is induced within the chamber. By designing this dynamic culture system, we ensured that the culture medium could flow evenly throughout the tissue construct.

Figure 4.

Figure 4

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Structural Differences Between Non‐Structured and Core–Shell Spheroidal Microtissue‐based Tissue Constructs Under Perfusion Culture Conditions. A,B) Schematic illustration of the custom‐designed perfusion culture system, along with actual images of the 3D‐printed PCL chamber (A) and its microstructure captured using SEM (B). C) Bright‐field images of the bottom regions of tissue constructs on days 0 and 7. D) Quantification of bottom area changes over time. Data are presented as mean ± SD (n = 3). E) Live/dead staining images of the top and bottom regions of tissue constructs on day 7. F) Hypoxia staining images of the bottom region of tissue constructs on day 7. G) Immunofluorescence staining of the bottom regions of tissue constructs on day 7 (4× and 10× images). H) H&E staining images of the top and bottom sections of tissue constructs on day 7. The first row displays representative 4× and 10× magnification images of non‐structured spheroidal microtissue‐based tissue, and the second row the corresponding images of core–shell spheroidal microtissue‐based tissue.

To compare tissue organization between non‐structured and core–shell spheroidal microtissues under perfusion culture at a flow rate of 50 µl min−1, 800 spheroids were loaded into the culture chamber and cultured for seven days. In this case, the initial volume of the spheroidal microtissue‐based tissue was 26.6 mm3. Figure 4C presents bright‐field images of the bottom regions of the tissue constructs on days 0 and 7, demonstrating significant contraction in the non‐structured spheroidal microtissue group, whereas the core–shell spheroidal microtissue group maintained its initial volume. This trend is further quantified in Figure 4D, where the bottom area of non‐structured spheroidal microtissue‐based tissues progressively decreased over time. In contrast, core–shell spheroidal microtissue‐based tissues retained a stable structure throughout the culture period. This difference in fusion resulted in significant differences in cell viability, as illustrated in Figure 4E, which shows live/dead staining images of the top and bottom surfaces of the two groups on day 7. There was no significant difference in the top region of the tissue; however, live/dead staining of the bottom region revealed substantial cell loss in the center of the non‐structured spheroidal microtissue‐based tissue, whereas the core–shell spheroidal microtissue‐based group maintained a higher level of viable cells throughout the construct. This difference in cell viability was further corroborated by hypoxic staining (Figure 4F). In the non‐structured spheroidal microtissue‐based tissue group, hypoxic regions were prominently observed in the bottom‐central areas, indicating insufficient oxygen supply due to excessive spheroid fusion and contraction. In contrast, the core‐spheroidal microtissue‐based tissue group exhibited a minimal hypoxic staining area.

Immunofluorescence staining was performed to examine the cellular distribution and internal structure of spheroidal microtissue‐based tissues (Figure 4G). The first row presents 4× and 10× magnification images of non‐structured spheroidal microtissue constructs, while the second row displays the corresponding images of core–shell spheroidal microtissue constructs. In the non‐structured spheroidal microtissue‐based tissues, the boundaries between spheroids became indistinct owing to fusion, and no intermicrotissue spaces were observed owing to the irregular proliferation of HepG2/C3A and EA.hy926 cells. In contrast, the core–shell spheroidal microtissue‐based tissue exhibited a more clearly defined spheroidal microtissue morphology. Remarkably, the spaces between spheroidal microtissues were also distinctly visible, and these spaces formed endothelialized channels (e.g., microvessels) owing to the EA.hy926 cells covering the spheroidal microtissue surfaces. These differences were more clearly observed in the higher‐magnification images in the second column.

Hematoxylin and eosin (H&E) staining was performed to assess the structural integrity of the tissue constructs after seven days of medium perfusion culture, as shown in Figure 4H. The non‐structured spheroidal microtissue‐based tissue exhibited significant volume contraction compared to the core–shell spheroidal microtissue‐based tissue, with extreme fusion and loss of the original spheroidal microtissue morphology. In particular, significant fusion of spheroidal microtissues was observed in the outer region of the tissue, where they were in direct contact with the culture medium. Consequently, the culture medium had likely flowed into the space formed between the culture chamber and the tissue due to contraction, rather than being evenly distributed throughout the tissue interior. This insufficient supply of culture medium may have contributed to cell death within the tissue. In contrast, the core–shell spheroidal microtissue‐based tissue maintained its spheroid morphology, consistent with the immunofluorescence staining results. Furthermore, moderate fusion occurred throughout the spheroidal microtissue‐based tissue construct. This moderate fusion allowed for the even flow of the culture medium during perfusion through spheroidal microtissue‐based tissue, enabling the fabrication and cultivation of tissue with high cell viability. This structural difference highlights the importance of endothelial cell‐covered spheroidal microtissues in tissue formation with microvascular networks under perfusion conditions.

2.5. Scaling Up Tissue Constructs by Controlling Medium Perfusion Rate

Core–shell structured spheroidal microtissues demonstrated excellent structural integrity, making them suitable for fabricating tissue constructs with integrated microvascular networks. However, these tissue constructs were still limited in volume. Therefore, as shown in Figure 5 , we leveraged the advantages of core–shell spheroidal microtissues to fabricate volumetric tissues. Using the perfusion culture system previously employed in Figure 4, we cultured a large‐volume construct containing 2400 spheroids, achieving a tissue volume of 79.3 mm3 (Figure 5A). At this time, the cell density within the tissue reaches 5 × 107 cells mL−1, resulting in a highly dense cellular structure (Note S1 and Figure S1, Supporting Information). We compared two different perfusion rates: maintaining the perfusion rate at 50 µL min−1, as shown in Figure 4, and increasing it to 300 µL min−1. On day 7, cell viability was assessed (Figure 5B). At a perfusion rate of 50 µL min−1, the tissue construct exhibited high cell viability only at the top surface, with significantly lower viability observed throughout the deeper regions. In contrast, tissue constructs cultured at 300 µL min−1 displayed excellent cell viability uniformly throughout the tissue structure (Figure 5C). This difference in cell viability notably affected the hepatic functionality of the embedded HepG2/C3A cells.

Figure 5.

Figure 5

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Culture Results for Volumetric Tissue and Investigation of the Influence of Medium Perfusion Rate. A) Photograph of volumetric tissue structure generated using core–shell spheroidal microtissues. B) Assessment of cell viability for volumetric tissue structures cultured at different medium perfusion rates. C) Distribution of live and dead cells along the height of tissue structures. D) Assessment of albumin, AAT, and urea production by tissue structures (n = 3). E) Comparison of ammonia elimination from the culture medium by core–shell spheroidal microtissue‐based volumetric tissue constructs according to perfusion rate. Data are presented as mean ± SD (n = 3). The asterisks indicate a significant difference between the 300 and 50 µL min−1 groups (p<0.05).

After assessing cell viability within the spheroidal microtissue‐based tissue construct, we conducted hepatic functional analysis mediated by parenchymal HepG2/C3A cells (Figure 5D). Albumin secretion, a critical hepatic parenchymal cell function involved in maintaining osmotic pressure and transporting fatty acids, hormones, and drugs, was quantified over 24 h periods under different perfusion rates. In tissue constructs cultured at two different perfusion rates, albumin secretion decreased from day 1 to day 3; however, it subsequently exhibited a common increasing trend up to day 7. However, throughout the entire culture period, tissue constructs cultured at a perfusion rate of 300 µL min−1 consistently produced higher levels of albumin compared to those cultured at 50 µL min−1. Alpha‐1 antitrypsin (AAT), another important liver‐produced protein, plays a crucial role in tissue protection and immune regulation through its anti‐elastase activity. The tissue constructs cultured at a flow rate of 300 µL min−1 exhibited a trend in AAT production similar to that observed in albumin secretion, with an initial decrease by day 3 followed by an increase up to day 7. In contrast, AAT production in tissue constructs cultured at 50 µL min−1 exhibited a decline between days 1 and 3 and continued to decrease through day 7, resulting in significantly lower AAT levels compared to constructs cultured at 300 µL min−1 at the end of the culture period. Additionally, quantitative analysis of urea production, a waste product derived from protein metabolism in hepatic parenchymal cells, was conducted. Tissue constructs cultured at a flow rate of 300 µL min−1 exhibited a decreasing trend in urea production from day 1 to day 3, followed by stabilization throughout the remainder of the culture period. In contrast, constructs cultured at 50 µL min−1 consistently produced lower levels of urea compared to those cultured at 300 µL min−1 over the entire culture duration. Furthermore, ammonia, a toxic byproduct generated during protein catabolism, is converted by hepatocytes into urea for excretion in the human body, underscoring the critical role of the liver in systemic homeostasis. The ammonia metabolism measured after day 1 and day 7 of culture confirmed that tissues cultured at 300 µL min−1 metabolized substantially higher amounts of ammonia than those cultured at 50 µL min−1 (Figure 5E). These phenomena of different cell viability and tissue functionality are presumed to result from differences in flow velocity and flow uniformity observed at the bottom surface of the culture chamber after culturing the tissues for 7 days under different perfusion rates (Note S2 and Movie S1, Supporting Information). These results strongly suggest that perfusion rate is crucial for determining cell viability and tissue functionality when producing volumetric tissues using bioprinted endothelial cell‐covered spheroidal microtissues.

3. Discussion

Developing bioengineered artificial organs to replace damaged tissues or organs is a critical challenge in medicine. Recently, genetically modified pig hearts were transplanted into patients.[ 35 , 36 ] While this represented a groundbreaking advancement in overcoming initial immune rejection, the patients died due to porcine cytomegalovirus infection or antibody‐mediated rejection. Given these challenges, biofabrication technology that utilizes human‐derived cells and biomaterials to create artificial tissues and organs is a promising therapeutic, regenerative, and replacement approach. Conventional biofabrication technologies have faced difficulties constructing tissues with high cellular density and microvascular networks, which are essential for tissue function, owing to their limited spatial resolution.[ 37 ] In this study, we successfully demonstrated a highly vascularized and high cell density tissue fabrication method by assembling bioprinted, endothelial cell‐covered spheroidal microtissues. Furthermore, we validated the feasibility of this approach for the production of large‐volume tissues. Consequently, the strategy for fabricating volumetric vascularized tissue presented in this study can be a promising approach for manufacturing engineered organ equivalents.

While this study represents a significant advancement, successful clinical translation requires consideration of vascular integration beyond the construct itself. Without proper vascular connections to the host circulatory system, implanted tissues risk ischemia and necrosis, undermining their therapeutic efficacy.[ 38 ] Therefore, incorporating vessel structures capable of connecting with the host's blood vessels becomes imperative. These external vessel connections facilitate nutrient exchange and tissue perfusion, promote tissue integration, and minimize the risk of graft rejection. Furthermore, the cells used in this study were cancerous or immortalized cell lines, differing significantly from native tissue cells in terms of proliferation, growth, and metabolism. Such disparities between the cellular composition of engineered tissues and native tissues could lead not only to differences in tissue organization processes during the in vitro assembly of microtissues but also serve as barriers to the clinical application of the fabricated tissues. Therefore, further research should explore the use of primary cells, such as patient‐derived hepatocytes and human umbilical vein endothelial cells, or stem cell‐derived cells, such as induced pluripotent stem cell‐derived hepatocytes and endothelial cells. Beyond mere tissue survival, they have profound implications for the clinical success and applicability of tissue‐engineered constructs, enabling complex tissue regeneration and organ replacement.

The successful translation of artificial tissue into viable clinical applications hinges upon several critical factors, one of which is its mechanical strength.[ 39 , 40 ] The ultimate goal of tissue engineering is to develop constructs that closely mimic native tissue, facilitating their seamless integration into the human body. Although our study primarily focused on the fabrication process and biological compatibility of the engineered tissue, we acknowledge the pivotal importance of mechanical strength for its eventual clinical utility. The mechanical strength of the artificial tissue produced in this study was not presented due to its low mechanical strength, which is lower than that of native tissue (less than 1 N at a strain of 0.005). This discrepancy underscores the need for further investigation into enhancing the mechanical characteristics of engineered constructs. One promising avenue for bolstering the mechanical properties of artificial tissues is through crosslinking techniques. While not employed in the current study, crosslinking agents such as genipin offer a straightforward approach to reinforcing the structural integrity of collagen‐based constructs.[ 41 ] Genipin‐mediated crosslinking improves the strength and stability of collagen matrices, rendering them more resilient to mechanical stress. Furthermore, incorporating fibrinogen into the bioink presents another avenue for enhancing mechanical strength.[ 42 ] By harnessing the enzymatic activity of thrombin, fibrinogen can be crosslinked within the bioink matrix, forming a robust fibrin network. This augmentation of the scaffold architecture holds promise for imparting greater mechanical stability to the fabricated tissue constructs. Furthermore, it is imperative to conduct comprehensive investigations into the mechanical properties of the fabricated tissue, including assessments of the tensile strength, compressive modulus, and shear resistance. Moreover, comparative analyses with their native tissue counterparts will provide invaluable insights into the efficacy of our fabrication approach. Additionally, exploring alternative crosslinking strategies and optimizing their parameters will be instrumental in further enhancing the mechanical characteristics of engineered tissue constructs.

The perfusion system used in this study operates based on hydrostatic pressure. This system is straightforward to construct, set up, and operate, as it does not require direct tubing or sealing of the culture chamber. Additionally, the system can include multichannel outlets, which help minimize blockages caused by dead cells or bubbles. However, this approach has some limitations. First, the liver comprises numerous functional units known as hepatic lobules. Each hepatic lobule exhibits a blood flow pattern in which blood supplied from the portal vein passes through a sinusoidal network, a microvascular structure situated between hepatocytes, and then drains into the central vein. In the present study, the volumetric hepatic tissue was fabricated by assembling spheroidal microtissues with a simple core–shell structure, resulting in blood flow characteristics that differ significantly from those observed in native hepatic lobules. Furthermore, controlling the pressure within the culture chamber is particularly challenging. The pressure is determined by the height of the culture medium, meaning that increasing the hydraulic pressure requires raising the height of the culture chamber,[ 43 , 44 ] making it difficult to elevate the system pressure. The system pressure also largely depends on the medium supply flow rate, which can fluctuate owing to variations in the peristaltic pump. Additionally, the height of the culture medium can be affected by the outlet conditions. Our chamber had eight outlets, and occasionally, these outlets were partially blocked by spheroidal microtissues, increasing the flow resistance and altering the media height. Pressure is crucial in mechanotransduction as it can influence cellular behavior. The interstitial pressure of the liver is 2–4 mm Hg,[ 45 , 46 , 47 ] which is higher than the pressure of the current open channel‐based system. Owing to the limitations of the current system in easily controlling pressure, it is challenging to replicate the native tissue environment accurately. As a result, the function and mechanical characteristics of the engineered tissue may differ from those of the actual tissue. Therefore, an advanced perfusion system is necessary to mimic the mechanical environment of native tissue better for effective tissue construct engineering.

Spheroid fusion has been extensively studied as a method for creating functional tissue constructs.[ 29 , 48 ] Some researchers have attempted to create vascularized tissue constructs using spheroids covered with an endothelial layer.[ 49 , 50 , 51 ] However, these efforts have often been limited regarding construct size and culture duration owing to the complete fusion of spheroids. In our study, generating a large quantity of uniformly endothelial‐covered spheroidal microtissues was essential for realizing our hypothesis, which was that the fusion of endothelial cell‐covered spheroidal microtissues fabricated using bioprinting‐based technology creates intermicrotissue spaces that function as a microvessel network. Among the spheroidal microtissues produced using our manufacturing method, over 95% were successfully covered with endothelial cells, providing a high‐quality, consistent foundation for vascularized tissue formation. Additionally, our system can produce 6000–8000 spheroidal microtissues in a single batch, a capacity currently limited by the precursor cartridge size. This production capacity can be easily scaled up by increasing the cartridge size, demonstrating the flexibility of the proposed method. Therefore, this 3D bioprinting‐based spheroidal microtissue production approach is ideal for producing engineered microtissues, such as endothelial cell‐covered or hepatic lobule‐like patterned microtissues,[ 34 ] and could support many tissue engineering applications.

The bioartificial liver (BAL) is a therapeutic approach designed to temporarily perform liver functions in patients, often serving as a bridge to transplantation or as supportive therapy during recovery.[ 52 ] BALs typically use hepatocytes to carry out liver functions. However, human hepatocytes are difficult to obtain, leading to the common use of porcine hepatocytes. For BALs, a critical challenge is maintaining a high density of functional hepatocytes over prolonged periods.[ 53 ] In our study, we developed vascularized, high‐density tissue constructs using endothelial cell‐covered spheroidal microtissues. By forming microvascular networks within the construct, these spheroidal microtissues enabled efficient nutrient and oxygen exchange, supporting large numbers of cells over time. This engineered tissue structure may offer a stable and biomimetic environment suitable for BAL applications. Our approach could potentially enhance the longevity and effectiveness of cell‐based BAL systems, creating a closer approximation of native liver tissue function.

4. Conclusion

In this study, we successfully assembled endothelial cell‐covered spheroidal microtissues to fabricate highly vascularized tissue constructs with high cell density. Our structured spheroidal microtissue fabrication method, leveraging preset bioprinting technology, enabled the uniform and high‐throughput production of endothelial cell‐covered spheroidal microtissues with precisely controlled core–shell structures. Compared to non‐structured spheroidal microtissues, endothelial cell‐covered spheroidal microtissues exhibited superior structural integrity, which proved crucial for their assembly into functional tissue structures. Tissue constructs based on non‐structured spheroidal microtissues, in which endothelial and parenchymal cells were uniformly mixed, exhibited irregular cellular arrangements and complete microtissue fusion, hindering the formation of functional tissues. In contrast, core–shell spheroidal microtissues successfully supported the formation of vascularized structures, particularly under dynamic culture conditions involving medium perfusion, which promoted the development of organized microvascular channel networks. Furthermore, we demonstrated that the perfusion rate of the culture medium significantly influenced cellular viability and tissue functionality within volumetric tissue constructs generated from core–shell spheroidal microtissues. These results underscore that the endothelial cell‐covered spheroidal microtissue‐based tissue fabrication strategy presented in this study can facilitate the development of large‐scale tissue constructs, serving as a foundational technology for creating clinically relevant tissue analogs.

Future research efforts should focus on optimizing strategies for vascular integration and external vessel connections, including the development of biomimetic scaffolds and refining surgical techniques. Long‐term preclinical studies are warranted to evaluate the safety, efficacy, and durability of vascularized tissue implants in relevant animal models. By addressing the need for external vessel connections, we can overcome key hurdles in tissue integration and pave the way for the development of functional implantable tissues with transformative potential in regenerative medicine.

5. Experimental Section

Cell Culture

HepG2/C3A cells (HB‐8065; ATCC, Washington, DC, USA) were used as a hepatocyte model, and EA.hy926 cells (CRL‐2922; ATCC) were used as an endothelial cell model for constructing spheroids. HepG2/C3A and EA.hy926 cells were cultured in Dulbecco's modified Eagle's medium (DMEM; 11885092, Gibco, Grand Island, NY, USA) supplemented with 10% (v/v) fetal bovine serum (16000044; Gibco) and 1% (v/v) penicillin‐streptomycin (15140122; Gibco) in an incubator at 37 °C with 5% CO2. The culture medium was exchanged every three days.

Preparation of Bioinks

Type 1 atelocollagen (MS collagen; MS Bio, Seongnam‐si, Gyeonggi‐do, Republic of Korea) was dissolved in 10−3 n HCl (pH 3.0) at a concentration of 4% for 3 days. The collagen solution was then mixed with 10× minimal essential medium (11430030; Gibco) and reconstitute buffer (2.2 g of NaHCO3, 4.77 g of HEPES, and 0.2 g of NaOH in 100 mL tertiary distilled water) at a volume ratio of 8:1:1. HepG2/C3A and EA.hy926 cells were dissociated with trypsin‐EDTA (25200056; Gibco) and mixed with neutralized collagen at concentrations of 4 × 107 cells mL−1 and 5 × 107 cells mL−1, respectively. Finally, the bubbles in the bioinks were removed using a centrifuge at 447 relative centrifugal force for 20 min. All procedures were conducted on ice or at 4 °C to prevent gelation of neutralized collagen.

Fabrication and Culture of Non‐Structured and Core–Shell Spheroidal Microtissues

Spheroidal microtissues were fabricated using the same preset extrusion bioprinting method combined with microfluidic emulsification as in the previous research.[ 34 ] Briefly, precursor cartridges with core–shell architecture were designed with 3D CAD software (Solidworks 2020; Dassault Systems, France) and fabricated with a 3D printer (ProJet 6000; 3D Systems, Circle Rock Hill, SC, USA). The fabricated precursor cartridges were then sterilized with 70% ethanol and UV light. The core and shell compartments of the precursor cartridge were filled with HepG2/C3A and EA.hy926 bioink, respectively. The precursor cartridge was transferred into a 3 mL syringe and extruded through the inner nozzle of a coaxial nozzle (NanoNC, Seoul, Republic of Korea) as a disperse phase with pneumatic pressure of 8–10 kPa. Mineral oil (330760; Sigma–Aldrich, St. Louis, MO, USA) with 2% v/v Span80 (86648; Sigma–Aldrich) was injected into the outer nozzle as a continuous phase. Finally, core–shell structured spheroidal microtissues were generated.

To fabricate non‐structured spheroidal microtissues, the HepG2/C3A and EA.hy926 bioinks were homogenously mixed at a volume ratio of 1:1.38 according to the volume ratio of the fabricated core–shell spheroidal microtissues (Note S3 and Figure S2, Supporting Information).

All spheroidal microtissues were fabricated using a commercial bioprinting system (3DX Printer; T&R Biofab, Siheung‐si, Gyeonggi‐do, Republic of Korea). After fabrication, microtissues were placed in a CO2 incubator for 1 h for gelation. To remove mineral oil from the culture dish, additional culture medium was added to the culture dish, and the upper layer of mineral oil was removed. The spheroids were cultured in DMEM in Petri dishes coated with 20 mg mL−1 poly‐HEMA (192066; Sigma–Aldrich). A poly‐HEMA coating was applied to prevent cell adhesion to the dish surface. Poly‐HEMA was dissolved in 95% ethanol at a concentration of 20 mg mL−1 and applied to the dish at a ratio of 20 µL mm−2. The coated dishes were then air‐dried inside a clean bench for at least 12 h. Following drying, the dishes were sterilized using ethylene oxide gas before use.

Static Culture

Static culture was performed to observe the tissue organization process of spheroidal microtissues without medium perfusion. Non‐structured and core–shell spheroidal microtissues were cultured in physical contact with each other on a 35 mm Petri dish coated with poly‐HEMA, using DMEM as the culture medium. Cultures were maintained at 37 °C in an incubator at 37 °C with 5% CO2, and the culture medium was replaced every 2 days.

Perfusion Culture

The perfusion culture system was designed to support large‐volume tissue constructs by ensuring uniform nutrients and oxygen distributions under medium flow conditions. To support nutrient and oxygen delivery within the tissue constructs, a perfusion culture system was designed and employed. The culture chamber was 3D‐printed using polycaprolactone and featured a cylindrical design with an octagonal cross‐section. It measured 15 mm in height and included eight bottom outlets, each measuring 400 µm in width and 200 µm in height. The chamber was 3D‐printed directly onto a 60 mm Petri dish, which was covered with a 3D‐printed polylactic acid. Silicone tubing (AY202354‐CP; Masterflex, Willy‐Brandt‐Allee, Gelsenkirchen, Germany) was connected to the cover, enabling the supply and circulation of the culture medium. A peristaltic pump (G100‐2J; Longer Precision Pump, Baoding City, Hebei Province, China) was used to maintain a continuous flow of culture medium. The perfusion system was installed in a CO2 incubator to provide the temperature and atmospheric conditions for cell growth.

Scanning Electron Microscope Image Acquisition

The 3D‐printed culture chamber was mounted on copper conductive tape and coated with platinum for 10 min. The samples were observed using field emission SEM (Nova NanoSEM 450; FEI, Waltham, MA, USA) with an accelerating voltage of 10 kV.

Calculation of Volume of Tissue

To calculate the volume of the tissue, spherical microtissues were placed in a dynamic culture chamber three times to measure the height of the tissue accumulated in the chamber. The height of the tissue formed by 800 microtissues was ≈2 mm, and the height of the tissue formed by 2400 microtissues was ≈6 mm. Then, the volume of a microtissue‐based tissue structure was calculated using the area of the dynamic culture chamber cross‐section, which is a regular octagon with a side length of 1.66 mm.

Analysis of Dimensions

The diameters and circularity of the spheroidal microtissues and the projected area of microtissue‐based tissue constructs were measured using an inverted microscope and analyzed with ImageJ software (NIH). The threshold function was applied to clearly define the bottom area of the tissue, allowing for the precise selection of the region of interest. Then, the area of samples was quantified within the threshold region. To eliminate edge effects, only spheroidal microtissues with their entire structure visible were included in the analysis.

Immunofluorescence Staining

HepG2/C3A cells in the spheroidal microtissues and spheroidal microtissue‐based tissue samples were stained with albumin antibody (ab207327; Abcam, Cambridge, UK) and hepatocyte nuclear factor 4 alpha (HNF4α) antibody (ab41898; Abcam), whereas EA.hy926 cells were stained with CD31 antibody (ab9498; Abcam). Nuclei were counterstained with Hoechst 33342 (H3570; Invitrogen, Waltham, MA, USA). To enable cross‐sectional observation, the tissues were rendered transparent using a tissue clearing/staining kit (V11325; Invitrogen). Samples were imaged using a confocal microscopy system (FV1200; Olympus, Tokyo, Japan).

Live/Dead and Hypoxia Staining

Live/dead staining was performed to assess cell viability within spheroidal microtissues and tissue constructs, whereas hypoxia staining was used to identify oxygen‐deficient regions within the tissue constructs. Live cells were stained with calcein‐AM (C1430; Invitrogen), and dead cells were stained with ethidium homodimer‐1 (E1169; Invitrogen). For hypoxia detection, samples were stained with Image‐iT Green Hypoxia Reagent (I14833; Invitrogen). All fluorescently stained samples were imaged using a confocal microscope under the appropriate fluorescence settings.

Paraffin Sectioning and H&E Staining

H&E staining was performed to visualize the histological structure and cellular organization within the tissue samples. Tissue samples were fixed with 4% paraformaldehyde at 4 °C for 12 h, dehydrated through a graded ethanol series (30%, 50%, 70%, 80%, 90%, and 100%), cleared in xylene, and embedded in paraffin. Tissue sections of 10 µm thickness were prepared using a microtome and mounted on glass slides. The sections were then deparaffinized in xylene, rehydrated through a reverse ethanol series, and stained with hematoxylin (HEMM‐OT; BioGnost, Zagreb, Croatia) and eosin (EOYA‐10‐OT; BioGnost). Images were acquired using a microscope (CKX53; Olympus) and a CMOS camera (eXcope T500; DIXI Science, Daejeon, Republic of Korea).

Enzyme‐Linked Immunosorbent Assay (ELISA), Urea and Ammonia Assays

To evaluate the hepatic function, the culture medium was collected on days 1, 3, 5, and 7 and stored at −80 °C until further analysis. Albumin concentration was measured using an albumin human ELISA kit (ab108788, Abcam), whereas urea concentration was determined using a urea assay kit (ab83362, Abcam). The levels of alpha‐1 antitrypsin were quantified using a human alpha‐1 antitrypsin ELISA kit (ab108799, Abcam). For analysis, the collected culture medium was thawed at 25 °C, and the optical density (O.D.) values were measured according to the manufacturer`s protocol. A standard curve was generated using the standard proteins provided by the manufacturer, and the concentration of albumin, urea, and AAT in the samples was determined by interpolating their O.D. values on the standard curve. For the ammonia assay, the culture medium was supplemented with 1 mM ammonium chloride (A9434, Sigma–Aldrich) on day 7 and incubated for 24 h. The culture medium was collected and diluted 1:10 before analysis. Dissolved ammonia in the culture medium was quantified using an ammonia assay kit (ab102509, Abcam). The amount of ammonia eliminated by the microtissue‐based tissue structure was calculated by subtracting the residual ammonia concentration in the sample from the initial 1 mM ammonia concentration in the untreated culture medium.

Statistical Analysis

All experiments were performed at least in triplicate unless otherwise stated. Data are presented as the mean ± SD. For statistical analysis, a t‐test was performed using Microsoft Excel software. Differences in results were considered significant at p<0.05.

Conflict of Interest

H.O., S.L., W.‐S.Y., J.‐H.S., and S.J. are employed by and shareholders of T&R Biofab. The remaining author declares no conflicts of interest.

Supporting information

Supporting Information

ADHM-15-0-s002.docx (197.3KB, docx)

Supplemental Movie 1

Download video file (5.9MB, mp4)

Acknowledgements

H.O. and J.‐H.K. contributed equally to this work. This research was supported by a National Research Foundation of Korea (NRF) grant funded by the Ministry of Education (Grant Number: NRF‐2017R1A6A1A03015562) and a Korean Fund for Regenerative Medicine (KFRM) grant funded by the Korean government (Ministry of Science and ICT, Ministry of Health & Welfare) (Grant Number: KFRM 21A0403L1).

Oh H., Kim J.‐H., Lee S., Shim J.‐H., Yun W.‐S., and Jin S., “Engineering Volumetric Tissue Analogs via Assembly of Endothelial Cell‐Covered Spheroidal Microtissues.” Adv. Healthcare Mater. 15, no. 2 (2026): e02418. 10.1002/adhm.202502418

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supporting Information

ADHM-15-0-s002.docx (197.3KB, docx)

Supplemental Movie 1

Download video file (5.9MB, mp4)

Data Availability Statement

The data that support the findings of this study are available on request from the corresponding author. The data are not publicly available due to privacy or ethical restrictions.

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