Heterogeneous liver tissues with multicellular crosstalk and bile duct morphogenesis through organoid bioprinting

doi:10.21203/rs.3.rs-2727920/v1

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Heterogeneous liver tissues with multicellular crosstalk and bile duct morphogenesis through organoid bioprinting

https://doi.org/10.21203/rs.3.rs-2727920/v1

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Liver is dynamic, heterogeneous, and each cell type acts in concert to regulate its function. Currently, heterogeneous liver tissues are typically built from single cells using bioprinting, making crosstalk between cells difficult. Therefore, in vitro morphogenesis is limited, and self-assembled biliary and blood vessels system are absent from manufactured liver tissues. The combination of bioprinting and organoid technique offers spatial and cellular control over three-dimensional (3D) organ tissue manufacturing, allowing to build liver tissues with self-assembled structure in vitro. We developed a high-throughput PDMS microwell platform (PMP) generating uniform and functional hepatic organoid building blocks (HOBBs) which displayed cellular crosstalk and self-assembled structure. For bioprinting process, we developed three-level temperature control system and new quadratic material, i.e., alginate-gelatin-collagen-laminin (AGCL) biomaterial, realizing reproducible construction of liver tissues with requisite cellular density. Under long-term differentiation, bioprinted liver tissues exhibited enhanced hepatobiliary function, intrahepatic bile duct networks and angiogenic potential. Heterogeneous tissues with coexistence of cholangiocytes, endothelial cells, and hepatocytes was constructed. The heterogeneous liver tissues with angiogenesis and bile duct component (HABs) provides a novel tool for morphogenesis study, liver regeneration, drug testing, and disease research.

Biological sciences/Biotechnology/Tissue engineering

Biological sciences/Biological techniques/Lab-on-a-chip

Liver functions as a metabolic hub and a critical regulator of blood homeostasis¹. Laboratory-made 3D liver tissues that fully maintain the function of human livers remain a major hope for liver regeneration, drug testing, tissue morphogenesis and disease development². 3D bioprinting, due to its ability to precisely organize multiple types of cells and biomaterials, has great potential to create key architectural elements that can direct the emergence of native functions and physiological structure in engineered organ³. To date, various bioprinting techniques are incorporated in creation of artificial heart^4,5, kidney^6,7, gastrointestinal tract^8,9, bladder¹⁰, and liver^11–13. Typically, dispersed hepatocyte cells are used to build liver tissues, and bioprinting technique has been successful in modelling cellular niche¹⁴. However, the liver tissues generally possess cell density one to two orders of magnitude lower, which disrupts interaction between cells, causing loss of phenotype and function. Tissue morphogenesis and development in vivo are predominantly determined by cell–cell crosstalk and microscale assembly, which also limited by single cell bioprinting method.

Bioprinting of cellular aggregates, such as spheroids^15,16, organoids^6,11, microtissues^17,18, which can mimic in vivo crosstalk and assembly of cells, offers viable strategy for constructing liver tissue with morphogenetic potential and mature function. Bernal et al¹¹ firstly created hepatic organoid-laden tissues that captured key liver functions and native-like polarization through volumetric bioprinting. Although their combination of bioprinting and organoid techniques offers new possibilities in liver regeneration, bioprinted liver tissues consist primarily of hepatic cells, making it difficult to simulate liver heterogeneity. Additionally, broader applications are subject to reproducibility of hepatic organoids.

Here, we developed a high-throughput PDMS microwell platform (PMP) capable of generating reproducible hepatic organoids with high oxygen permeability and soft mechanical property. HepaRG cells are proliferative hepatoblast-like cells, possessing stable bidirectional differentiation potential towards hepatocytes and cholangiocytes and facilitating generation of reproducible hepatobiliary organoids¹⁹. Through the bud stage, hepatoblasts and endothelial cells interact to promote liver development²⁰. Prior to functional blood vessel, vasculogenic endothelial cells are more critical for early stages of liver organogenesis²¹. We co-cultured HepaRG cells and HUVECs using PMPs to develop HOBBs. Changed expressions of VEGFA, Stab2, HGF, etc indicated the crosstalk between HepaRG cells and HUVECs like hepatoblasts-endothelial cells. HUVECs lined the nascent HOBBs and physically interacted with HepaRG cells, forming a self-assembled structure. Microextrusion bioprinting has been extensively researched and is considered the most promising method for manufacturing 3D tissues in a realistic timeframe²². However, gravity causes an inhomogeneous distribution of organoids during printing process. We developed three-level temperature controller system that prevented settling of organoids, allowing to print uniform liver tissues with requisite cell density. Based on uniform HOBBs and stable bioprinting system, we produced liver tissues with little individual and batch variation in a high-throughput manner. Biomaterial is critical for microextrusion-based organoid bioprinting process and post-printing neotissue formation. To promote organoids viability and hepatobiliary function, we developed new quadratic material, i.e., AGCL biomaterial. HOBBs were intact and highly viable after printing, and showed primary intrahepatic bile duct (IHBD) morphological structure and relatively mature hepatobiliary function in bioprinted constructs with media inducted. Also, bile duct and blood vessel development-related signaling pathway was upregulated in RNA-seq results. The study preliminarily explored the construction of liver tissues contained blood vessel and bile duct components in vitro, and provided novel systematic strategy for liver regeneration.

The development of PMPs

We aimed to build a high-throughput organoid culture platform (500 microwells in 1cm²), and Fig. 1a summarized the overall strategy. Briefly, we developed PMPs by transcribed silicon mold structures accurately. As the microwell of PMP had a special inverted pyramid structure (the side length of tightly arranged microwells is ~ 450 µm and the depth is ~ 400 µm, as shown in Fig. 1b and Fig. 1c), which was difficult to broadly transcribe in PDMS using conventional molding machine. We achieved highly accuracy by employing three-step transcription method (see Experimental Procedures). Further, PMPs could be punched to fit with any type of multi-well culture plates, flasks, and petri dishes. Hepatic organoids were generated from only HepaRG cells for PMP characterization. The size of organoids could be controlled by adjusting input cell density (as shown in Fig. 1d). 90, 110, 130, and 150 µm diameter organoids were generated by seeding 400, 600, 800, and 1000 cells in each microwell in day4, respectively. Organoids' controllable size allows more flexibility in optimizing printing parameters.

Compared to low attachment plates, organoids generated by PMPs were consistent in size and shape, uniform within and between independent experiments (Fig. 2a). Uniform generation of organoids made it possible to construct reproducible liver tissues. Regulated changes in reactive oxygen species (ROS) activities are important to maintain normal development of hepatoblasts²³. As shown in Fig. 2b, intracellular ROS level of hepatic organoids was significantly enhanced in PMP groups compared to Aggrewell™ groups. Intracellular ROS activities are influenced by changes in environmental oxygen concentration²⁴. The high oxygen permeability of PDMS makes it ideal for use in hepatic organoid culture. Increasing evidence suggested that liver homeostasis and function depend on mechanical properties (e.g., stiffness and shear stress) ^25,26. We optimized curing temperature and time of PDMS to balance transcription accuracy and mechanical properties. The stiffness of PMPs was 3.3 ± 0.07 MPa. Aggrewell™ plates were stiffer than PMPs by three orders of magnitude (Fig. 2c). Hepatic organoids cultured in PMPs with lower stiffness and high oxygen permeability demonstrated higher cell activity, upregulated ALB and MRP2 gene expression than Aggrewell™ groups, suggesting that PMPs are more suitable for hepatic organoids generation (Fig. 2d, e).

Self-assembly and multicellular crosstalk of HOBBs

Throughout the development of the liver, multicellular crosstalk occurs. Hepatoblasts and endothelial cells interact during liver bud development²⁷. Emergence of liver bud requires inductive signals from endothelial cells. Prior to liver bud expansion, endothelial cells line the nascent sinusoids and physically interact with the hepatoblasts²¹. Hepatoblasts were also identified as a positive stimulator of sinusoid morphogenesis and maturation²⁸. For the development of liver, crosstalk between hepatoblasts and endothelial cells is essential. Recent advances in developing in vitro liver tissue models show that function of hepatoblasts is altered when co-culturing with endothelial cells^13,29,30. Although HUVECs are not native to liver, many have used them due to their ease of access and established protocols³¹. To generate HOBBs, we co-cultured HepaRG cells and HUVECs with PMPs upon mixture ratio (Supplementary Fig. 1). Comparing cell activity and gene expression of each group, we selected the group that had a cell ratio of HepaRG cells to HUVECs of 2:1 (Supplementary Fig. 1).

By comparing hepatic organoids and HOBBs in day1 (Fig. 3a, Supplementary Fig. 2a), it was found that HUVECs accelerated organoids formation, implying interaction and crosstalk between HepaRG cells and HUVECs. Cell aggregation to form organoids is a multi-step process characterized by successive changes in cell morphology and gene expression, including regulation of actin cytoskeleton, cell-cell assembly, remodeling of extracellular matrix, and cell-matrix adhesion³². Genes regulating actin cytoskeleton signaling pathways such as MYL6, ZYX, ARPC4, TMBS4X, CFL1, PFN1, ACTG1, and ACTGB were upregulated in HOBB group versus 2D group, proving that cells underwent morphological changes and regulation of actin cytoskeleton (Fig. 3b, Supplementary Fig. 2b). Cell–cell adhesion through molecules such as ICAM1, JAM1, JAM2 and Nectin4 genes was required for organoids formation, their upregulation suggesting tight assembly of cells in HOBBs group (Fig. 3b). CLDNs are major structural components of tight junctions (TJs) and regarded as cellular backbone, tighten the paracellular cleft of epithelium and endothelia against unwanted passage of solutes. Epithelial and endothelial cell-cell junction genes CLDN3, CLDN 4, CLDN 5 and CLDN 14 expression of HOBBs were upregulated (Fig. 3b). The above change indicated HUVECs and HepaRG cells within HOBBs were tightly connected, establishing the basis for crosstalk between them. Extracellular matrix (ECM) secretion played a significant role in organoid morphology³³. HOBBs exhibited enhanced matrix secretion and higher expression levels of COL9A3, COL9A2, COL6A3, and LAMB3, and the corresponding integrins ITGA2, ITGA5, ITGB5 and other ECM receptors MMP9 were also upregulated (Fig. 3b). According to these findings, HOBBs assembled dynamically, remodeled the extracellular matrix, formed tight cell-cell and cell-matrix connections. HOBBs possessed more bionic niches than 2D environment.

Constant confocal wide-field imaging and fluorescence-based quantitative assessment of collected HOBBs generated from multiple stages (day 0–4) revealed the migration of HUVECs-GFP (Fig. 3c). HOBBs’ fluorescence intensity decreased inside and increased at the edge. In order to elucidate HepaRG cells and HUVECs locations within HOBBs, we performed co-immunostaining with ALB and CD31. ALB⁺ HepaRG cells were found within HOBBs, and CD31⁺ HUVECs surrounded HOBBs (Fig. 3d). Gene set enrichment analysis (GSEA) of RNAseq data showed upregulation of VEGF signaling pathway of HOBB groups (Fig. 3e). Due to the VEGF signaling pathway regulates endothelial cell migration, its upregulation explained HUVEC migration-associated gene changes.

As mentioned above, we used HepaRG cells, proliferative hepatoblast-like cells, and HUVECs to generate HOBBs. It is essential for the crosstalk between hepatoblasts and endothelial cells to occur during liver development, as the secretion of cytokines and growth factors is influenced by each other. The upregulated expression of VEGFR2, vWF, CD31, LYVE1, HGF, Stab2, CD32b and VEGFA which participate in hepatoblasts-endothelial cells crosstalk, was observed in HOBB groups (Fig. 3f). Hepatoblasts work as a positive stimulator of endothelial cells morphogenesis and maturation, in which HGF together with VEGF play a decisive role^21,34. HGF and VEGF significantly promote expression of sinusoidal endothelial cell markers, such as CD32b, LYVE1, and Stab2, and suppress the expression of angioblast cell markers, such as VEGFR2, VWF, and CD31^20,28. We hypothesize that the crosstalk between HepaRG cells and HUVECs was similar to that between hepatoblasts and endothelial cells at liver bud stage, and HOBBs developed dynamically.

Based on qRT-PCR results, HOBB groups showed higher ALB, MRP2, BSEP expression and reduced AFP expression relative to 2D groups (Fig. 3g). Biosynthesis of amino acids, glycolysis-gluconeogenesis signaling pathway enriched by GSEA were upregulated in HOBB groups (Supplementary Fig. 2c). HOBBs had more mature liver function than 2D cells. SOX9, CK19, HEY1, and HES1, landmark genes expressed during bile duct development, were upregulated, suggesting the HOBBs facilitated cholangiocytes differentiation (Fig. 3h). So far, we developed reproducible building blocks with primary hepatobiliary function and multicellular crosstalk for manufacturing large-scale liver tissues.

Reproducible bioprinting of HABs

ECM is complex macromolecular structural network surrounding cells, which is not only a physical scaffold, but also a crucial modulator of biologic processes including cell attachment, migration, differentiation, repair, and development³⁵. Recent studies indicated that ECM modulates hepatic development, regeneration, and even maintenance of the normal architecture and differentiated state³⁶. Collagen Ⅰ is one of the most abundant ECM components supporting hepatocytes in liver cords³⁷. Laminin mainly locates near bile ducts, and is sufficient as a component of basal lamina for the commitment of bipotential liver progenitors to cholangiocytes. Especially, α5-containing laminin is necessary for formation of mature bile duct structures, and laminin 511 improves differentiation of cholangiocytes³⁸. Moreover, collagen I and laminin 511 is coincident with angiogenesis, and commonly used as biomaterials for angiogenesis assay^39–41. To promote bile duct morphogenesis and angiogenesis, the liver tissues were built using gelatin-alginate-based bioink (AG) containing collagen I and laminin 511. We added 0.5 mg/ml collagen I and 10 µg/ml laminin 511 to gelatin-alginate-based bioink, developing new quadratic bioink, i.e., AGCL bioink, as shown in Fig. 4a. With AGCL bioink, the 3D-printed constructs had a clear and stable structure with interconnected channels and networks, and the filaments were uniform and smooth (Fig. 4a). SEM image of bioink demonstrated that it was porous, penetrating and scattered with protein fibers, facilitating cell metabolism, migration, and adhesion (Fig. 4b). Integrins constitute a major group of receptors for extracellular matrix components⁴². Currently collagen-binding I domain-containing integrins are known, namely ITGA2 and ITGB1⁴². Laminin 511 binding integrins are ITGA6 and ITGB1⁴¹. According to qRT-PCR results, the AGCL groups expressed higher levels of ITGA2, ITGB1, and ITGA6 than AG groups in day 14 (Supplementary Fig. 5), indicating cells were interconnected with collagen Ⅰ and laminin 511 in HABs.

Different from nozzle-free bioprinting technologies (e.g., stereolithography, laser-direct-writing), the microextrusion approach relies on a microneedle to deliver bioinks and print constructions, where the printability of bioink plays a crucial role⁴³. Printability is highly dependent on the rheological properties of bioink, so it is necessary to test its rheological behaviour. Prior to gelation, both AG and AGCL bioink showed a typical fluid-like behaviour (G′′ > G′) (Fig. 4c). Upon cooling, G′ for both solutions increased rapidly and eventually crosses over G′′ showing characteristics of a gel-like structure (Fig. 4c). AGCL bioink's gelation temperature (where G′ and G′′ cross over) was 4°C higher than AG bioink at 27°C, as shown in Fig. 4c. This suggested that printing of AGCL bioink should be conducted at higher temperature to ensure the proper gelation statuses. A print temperature of 27°C had no significant effect on cell activity. Long-term mechanical stability of bioink is required for high-throughput 3D bioprinting. We evaluated the changes in G' and G'' of AGCL bioink through time sweep tests, and it maintained stable viscoelasticity within 30 minutes. In particular, the G' for AGCL was 22.37 ± 0.35 Pa and G'' was 9.15 ± 0.18 Pa with a smoother curve (Fig. 4d). Another key characteristic of bioink for microextrusion printing is shear-thinning. As shown in Fig. 4e, shear force continuously decreased AGCL's viscosity, and it had good shear thinning properties which can prevent clogging and decreasing shear stress from damaging cells. With appropriate printability, AGCL bioink can be used in microextrusion bioprinting and layer-by-layer fabrication.

Creating reproducible liver tissue models requires not only uniform building blocks, but also stable printing process. HOBBs are prone to settling down during long-time printing, so we developed three-level temperature control system which possess precise temperature control (± 0.1℃) to prevent the settlement. In brief, three parts controlled the crosslinking status of bioink: syringe, needle, and substrate. The bioink in syringe was kept in a gel state to prevent the settlement of HOBBs, and microfilaments extruded from nozzle were kept proper crosslinked to print a clear outline and structure. Then, the printer's platform temperature was controlled to maintain a low temperature for supporting bioprinted constructions. HOBBs within constructions were uniformly distributed, allowing for reproducible liver tissue fabrication.

We printed larger HOBBs and found high cell viability after printing (Fig. 4f). This indicates our printing system (including bioink, printing process, and printing parameters) was well adaptable for HOBBs. During culture process, HOBBs within constructions gradually grew larger, and by day 14, their size was averaging 300µm (Supplementary Fig. 6). Due to excessive size and bioink blockage, cells cannot exchange substances resulting in advanced necrosis (Fig. 4f and Supplementary Fig. 7a). The maximum distance between cells and capillaries supplied nutrients and oxygen is 200 µm in vivo. Taking into account growth pattern of HOBBs in scaffolds, we eventually selected HOBBs with 90-µm initial size, and the bioprinting density is 1.06 ± 0.15×10⁷cells/ml (15000 organoids/ml) (Supplementary Fig. 3, 4). The viability of HOBBs remained high until day 14 without necrosis or apoptosis, satisfying in vitro culture requirement of large-scale liver tissues for long period of time (Fig. 4f).

Differentiation and liver function validation of HABs

HepRG cells, exhibiting differentiated cells merging with cholangiocyte and hepatocyte phenotype under DMSO treatment, are frequently described⁴⁴. The bipotent cell line HepaRG cells constitutes an efficient surrogate of liver function, yet its differentiated status relies on high concentrations of DMSO and is time-consuming, which may compromise the study of drug metabolism and limit the applicability of hepatic model⁴⁵. Herein, in this work, only a concentration of 0.5% DMSO was required to induce the differentiation of HepaRG cells within 14 days.

HOBBs in scaffolds expanded gradually based on light microscopic examination at different stages, as shown in Supplementary Fig. 6 and Fig. 5a. Cells started sprouting from HOBBs into scaffolds on day 7, and the sprouting cells formed a network by day 14 (Fig. 5a). The cell density had reached 3.33×10⁷/ml at day14 (Fig. 5b). According to SEM image, there is a clear difference between matrix with and without HOBBs, indicating that the HOBBs remodeled the matrix (Fig. 5c). Different morphologies indicated different cell types as shown in Fig. 5a. To determine types of cells were sprouting and aggregating in scaffolds, we performed ALB/CD31 co-immunostaining. Results showed that ALB⁺/CD31⁺ cells constituted the aggregated organoids in HABs, but neither ALB nor CD31 were staining cells sprouted in scaffolds (as shown by arrows in Fig. 5e). PAS staining is a tool to mark carbohydrates, and commonly used to evaluate glycogen storage and function of mature hepatocytes⁴⁶. As expected, HABs exhibited high levels of glycogen storage, acquired a characteristic hepatocyte-like morphology (Fig. 5f). Cryosectioning immunostains further revealed that HABs expressed higher mature liver markers, including ALB, CYP3A4 and MRP2, in comparison with HOBB groups (Fig. 5d).

Based on KEGG pathway analysis of liver function in HAB versus HOBB groups, we found enriched Hippo signaling pathways, NOD-like receptor signaling pathways, FoxO signaling pathways, and thyroid hormone signaling pathways were significantly upregulated, which are important for maintaining hepatocyte function and essential for determining their fate (Fig. 5i). GSEA revealed that the signaling pathway involved in gluconeogenesis, drug metabolism, fatty acid metabolism, biosynthesis of unsaturated fatty acids, N_glycan biosynthesis and histidine metabolism were enhanced in HAB versus HOBB groups, and the expression of related genes was also upregulated (Supplementary Fig. 7b, c). CD31 and VWF levels were reduced in HAB group, while Stab2 and F8 were upregulated, identifying characteristics of convergence to LSECs (Fig. 5g). Gluconeogenesis, urea synthesis, and fatty acid degradation related genes involved in the tricarboxylic acid cycle (TCA) were enhanced in HAB groups compared to HOBB groups. Clearly, the HABs were in constant dynamic change and displayed relatively mature liver function.

HABs with bile duct morphogenesis

HepaRG cells can differentiate into both hepatocytes and cholangiocytes. As mentioned previously, organoids aggregated within HABs were hepatocytes and HUVECs. HOBBs were composed of HepaRG cells and HUVECs, supporting the hypothesis that the cells spreading within HABs were cholangiocytes. We performed CK19/VE-Cad co-immunostaining and identified the spread cells as CK19⁺/VE-Cad⁻ cholangiocytes (Fig. 6a). Cryosectioning immunostaining revealed the spread cells within HABs expressed cholangiocytes markers as well, including SOX9, Ep-CAM and Cilia (Fig. 6c). Occasionally, cholangiocytes formed tubes with a central lumen as shown in red box (Fig. 6a). Further, fluorescent bile salt was transported by functional cholangiocytes incubated with cholyl-lysylfluorescein (CLF) (Fig. 6b). RNA-seq results revealed upregulation of ABCC4, ABCB1, PRKACB, PRKACB, ADCY3, ATP1B1, ATP1A1, SLC4A4 genes associated with bile acid salt transport and secretion (Fig. 6f). It was suggested that cholangiocytes within HABs had potential to form functional bile ducts. Biliary system is made up of intrahepatic bile duct (IHBD), extrahepatic bile duct (EHBD) and the gallbladder. HAB groups had noticeably higher expression level in intrahepatic bile duct (IHBD) marker genes than extrahepatic bile duct (EHBD) and gallbladder (Fig. 6d). The cholangiocytes interwoven with the hepatocytes were closer to the IHBD cells. FGF signal pathway, TGF-β signal pathway, Notch signal pathway and Wnt signal pathway were main pathways to regulate cholangiocytes differentiation and bile duct morphogenesis in vivo. In HAB groups, their signature genes CK19, SOX9, HES1, HES4, Notch1, BMP4, Wnt5, HNF1β, β-catenin, Jagged2, Smad4, TGFβ2, TGFβ3 and TGFβR2 were altered (Fig. 6h). SOX9, CK19, Hes1, Hes4 and Notch1 were also upregulated in the HAB groups based on qRT-PCR results (Fig. 6e). These data suggested that appropriate signaling pathways were activated to be involved in biliary differentiation. Altogether, part of HepaRG cells differentiated into functional cholangiocytes successfully.

Bile duct development can be divided into two steps, i.e., cell fate decision and morphogenesis²⁷. As shown in Fig. 6a and c, part of HepaRG cells differentiated into cholangiocytes successfully. Then, cholangiocytes form ductal plates, and reorganize into bile duct tubules, which depend matrix remodeling and cellular self-assembly⁴⁷. Gene Ontology (GO) enrichment analysis displayed that HAB groups upregulated in cell assembly and matrix organization related signaling pathway, including occluding junction, apical junction assembly, adherens junction assembly, focal adhesion assembly, extracellular matrix organization, and tube morphogenesis and branching related signaling pathways, including epithelial cell migration, regulation of cell morphogenesis, morphogenesis of a branching epithelium, morphogenesis of an epithelial sheet, epithelial tube morphogenesis, branching morphogenesis of an epithelial tube compared to HOBB groups (Fig. 6g). Based on GO enrichment analysis, cholangiocytes in HABs were continually assembling and remodeling extracellular matrix, indicating HABs’ potential for bile duct morphogenesis.

As IHBD develop from bipotent hepatoblasts around the portal vein, angiogenic signaling and crosstalk between them are vital to its development⁴⁸. Gene Ontology (GO) enrichment analysis showed that HAB groups were positively associated with angiogenesis (Fig. 7b). VEGF/VEGFR-2 signaling pathway links bile duct morphogenesis to angiogenesis⁴⁹. VEGFA, and other angiogenic signals in VEGF/VEGFR-2 signaling pathway, such as FAK, ERK, are enhanced expressed in HAB groups (Fig. 7c). Ang-1 represents a group of tyrosine kinase receptor ligands of VEGFA, which primarily play a role in developmental vascular remodeling and angiogenesis, also upregulated (Fig. 7c). The potential of HABs to form blood vessels was demonstrated. Enriched PPI network, which consisted of significantly enriched angiogenesis and biliary development signaling pathways, further evidenced crosstalk between bile duct morphogenesis and angiogenesis in HABs (Fig. 7d).

The first phase of the study involved developing PMPs capable of generating highly uniform and functional organoids. PMPs demonstrated higher oxygen permeability and softer mechanical properties than Aggrewell™ plates, which can promote hepatic organoid activity. As the emergence of liver bud requires inductive signals from endothelial cells and hepatoblasts, we selected hepatoblast-like cells, i.e., HepaRG cells, and HUVECs to generate bioprinting building blocks. HepaRG cells and HUVECs comprised HOBBs were distributed in an ordered manner, with HUVECs at the edges of HOBBs and HepaRG cells inside HOBBs, and similar hepatoblasts-endothelial cells crosstalk were showed between them. Self-assembled HOBBs with primary biological functions and multicellular crosstalk were derived, which paved the way for reproducible manufacture of heterogeneous liver tissues. Besides, PMPs provides an ideal tool on building blocks generation for other complex organs. In combination with advanced biofabrication technologies, such as organ-on-a-chip, 3D bioprinting, etc, more complicated mimicking organs can easily be made.

For bioprinting process, we developed three-level temperature control system and new AGCL biomaterial, realizing stable construction of liver tissues with requisite cellular density. Based on uniform HOBBs and robust bioprinting system, we produced reproducible large-scale liver tissues. With long-term differentiation process, HABs exhibited enhanced hepatobiliary function, intrahepatic bile duct networks and angiogenic potential. Heterogeneous tissues with coexistence of cholangiocytes, endothelial cells, and hepatocytes were formed. The results shed insights on the morphogenesis process of hepatobiliary architecture with branching networks via exogenous chemical induction. The heterogeneous hepatic tissues with multicellular crosstalk and bile duct morphogenesis provided novel tool for drug testing, disease research, and liver regeneration.

2D HepaRG cells culture

Undifferentiated HepaRG cells (Sigma-Aldrich, HPRGC1) were cultured in growth medium, which was composed of DMEM (Gibco,119600440) supplemented with 10% FBS (BI, 04-001-1A), 1% GlutaMAX™ supplement (Gibco, 35050061), 1% MEM NEAA solution (Gibco, 11140050), 0.05% insulin (Sigma, I9278), and 5×10^− 5 M hydrocortisone sodium succinate (Changzhou Siyao Pharm, China). HepaRG cells were incubated in humidified air containing 5% CO₂ at 37°C. When 85%-90% confluent, HepaRG cells were dissociated and passaged with trypsin/EDTA (Gibco, 25200056).

2D HUVECs culture and lentiviral transfection

Human umbilical vein endothelial cells (HUVECs) were purchased from NICR and cultured in DMEM (Gibco, 119600440) supplemented with 10% (v/v) FBS (BI, 04-001-1A), 0.1% insulin (Sigma, I9278), and 5ng/ml VEGF (100-20-500, Peprotech). Previously described lentivirus-mediated GFP kit (REVG1001, Genechem) was used to transfect HUVECs⁵⁰. HUVECs-GFP were also maintained in HUVECs growth medium and passaged when 70–80% confluence had been reached, changing the medium daily.

PMP fabrication and mechanical testing

PMPs for HOBB generation were fabricated based on a modified three-step protocol. In the first step, silicon wafers with microwells were used as templates to manufacture PMMA molds. The PMMA mold was placed on silicon wafers and then pressed by Multi-purpose Press (Model WY-30T form Dongguan Weiying Machinery (Equipment) Co., Ltd.), held at 150℃ for 3 minutes. Sylgard 184 (Dow Corning, USA) base A and curing agent B were then mixed in a weight ratio of 10:1. The PMMA mold was filled with mixture and kept at 70°C for 80 minutes after degassing to remove trapped bubbles. As the final step, PMPs were removed from PMMA mold and punched to fit a cell culture plate for generation of HOBBs. By using an Agilent Nano Indenter (G200, Keysight, USA) in CSM mode with a Berkovich tip at ambient temperature, the mechanical properties of PMP and Aggrewell™ plate (34415, Stemcell) were evaluated. Three samples were tested for each group(n = 3).

HOBB formation and culture

PMPs were first sterilized by autoclave before use. After fitting in 24-well plate, PMPs were rinsed with 1% w/v Pluronic® F-127 (Sigma-Aldrich, P2443), centrifuged for 5 min at 2000g and incubated at room temperature for 2h to prevent cell adhesion and promote efficient HOBBs formation. Subsequently, cell suspensions of HepaRG cells and HUVECs were prepared with a 2:1 ratio and seeded at 1000 cells/microwell, 800 cells/microwell, 600 cells/microwell, and 400 cells/microwell, respectively. PMPs were centrifuged for 5 min at 100g to ensure all cells were captured in microwell and then incubated at 37°C in 5% CO₂ with medium replenished every day. HOBBs were cultured in HepaRG cell growth medium. HOBBs were formed in ultra-low attachment plates (EFL-SP201, China) and Aggrewell™ 400 as described in protocol.

Determination of intracellular reactive oxygen species (ROS)

Based on the intracellular peroxide-dependent oxidation of DCFH-DA to form fluorescent compound 2',7'-dichlorofluorescein (DCF), intracellular reactive oxygen species (ROS) were measured. HOBBs were incubated for 30 minutes at 37°C with 20 mM DCFH-DA. HOBBs were then washed twice with PBS while fluorescence intensity was measured using a Laser scanning confocal microscope (Nikon, Z2).

Live/Dead staining

Fluorescent live/dead staining was applied to evaluate cell viability. Briefly, HOBBs or HABs were washed with PBS and incubated with 1 µM Calcein-AM (Sigma-Aldrich,17783) and 2 µM propidium iodide (Sigma-Aldrich, P4170) to stain live(green) and dead cells (red), respectively, then imaged by Laser scanning confocal microscopy (Nikon, Z2).

Bioink preparation and characterization

Gelatin (Sigma-Aldrich, G1890) and Alginate acid sodium salt powder (Sigma-Aldrich, A0682) were dissolved in 0.9% sodium chloride to prepare solutions as previously described⁵¹. The AGCL bioink was obtained by mixing Alginate, Gelatin, Collagen I (Advanced Biomatrix, 5056), and Laminin 511 (Biolamina, LN511-0502) solutions at 1% (w/v), 5% (w/v), 0.5 mg/ml, and 10 mg/ml concentrations and neutralizing bioink with 0.1mM NaOH to pH 7.0. HOBBs were collected by centrifugation at 100g for 3 min and resuspended by AGCL bioink reaching a final concentration of 15000 organoids/ml.

Rheological measurements were performed with a rotational rheometer (MCR302, Anton Paar, Germany) using a 25mm-diameter cone-plate with a 2° cone and 99-µm truncation gap as previously reported⁵¹. A temperature sweep test was carried out from 5°C to 40°C with the constant frequency of 1 HZ and 1% strain to obtain proper crosslinking temperatures. To determine the shear-thinning properties of AGCL bioink, controlled shear rate (CSR) mode with shear rate ranging from 0.01 s^− 1 to 100 s^− 1 under a controlled frequency of 1 rad/s was set and the viscosity was recorded every 5 s. The oscillatory mode at a 0.1% strain and 1.5Hz frequency was used to determine the rheological properties of AGCL bioink under optimized printing conditions. AGCL bioink was incubated at 37°C for 5 min and then loaded onto the platform. Following loading, the sample was cooled to gelation temperature and then tested for 35 minutes, recording the storage modulus (G’) and loss modulus (G’’).

AGCL bioink was prepared and lyophilized as previously described⁵¹. The samples were sprayed with a thin gold layer and examined under a field emission scanning electron microscope (JSM-7800F, JEOL, Japan).

3D bioprinting and culture of HABs

A commercial microextrusion-based 3D bioprinter (Regenovo Biotechnology Co., Ltd, Bio-architect X) whose temperature could be set between − 5℃ and 150℃ were used to fabricate HOBB-laden scaffolds. Regenovo software was used to design print structure and to regulate print parameters in real time. The temperature of sterile chamber, detachable nozzle and movable base plate was set at 24, 27, and 10℃, respectively, which is to facilitate instantaneous crosslinking of bioink and 3D construct formation. A six-layered grid scaffold (10mm×10mm) was printed with filament distances of 1.6 mm and layer distances of 150 µm. By using the extrusion-based printer, 3ml of HOBB-laden bioink was extruded at 0.14 MPa, 6mm/min through a 23G needle. Upon further crosslinking with 200 mM CaCl₂, the constructs were washed with PBS (02-024-1ACS, BI), and immediately transferred to differentiation medium (HepaRG cells growth medium supplemented with 0.5% DMSO) and cultured at 37℃ in 5% CO₂. To assess the effect of the manufacturing process on cell viability, scaffold survival was evaluated after printing. Cultured medium was renewed every 1 ~ 2 days.

Flow Cytometry of Apoptosis of HABs

After rinsing the scaffold with PBS, calcium ions in HABs were chelated for 5–10 minutes with a de-cross-linking solution (150 mM sodium chloride containing 55 mM sodium citrate and 20 mM EDTA). The organoids within HABs were collected by centrifugation for further assessment. Collected organoids were dissociated to single cell by trypsin/EDTA at 37℃ for 5min. The apoptosis of cells was demined by flow cytometry using a dead cell apoptosis kit with Annexin V Alexa Fluor™ 488 and propidium iodide (C1062S, Beyotime). Binding buffer containing 10 µl Annexin V and 5 µl PI was added to samples for 1h at 4℃. Novoexpress flow cytometry (BD Biosciences, San Jose, CA, USA) and Novocyte software were used to analyze the apoptosis of cells.

qRT-PCR Analysis

Trnzol (DP424, TIANGEN) was used to extract total RNA. The reverse transcription was performed using a cDNA synthesis kit (KR116, TIANGEN) following manufacturer's instructions. Maxima SYBR Green qPCR master mix (FP205, TIANGEN) was used with a real-time PCR detection platform (Bio-Rad, CFX96). Data were analyzed using BioRad CFX manager. Each sample was analyzed in triplicate. The housekeeping gene, glyceraldehyde3-phophate dehydrogenase (GAPDH), was used as an internal control for normalization. Primers used are listed in Supplementary Table S1.

Cryosectioning & Immunofluorescent staining

4% paraformaldehyde (PFA) (P1110, Solarbio) was added to HOBBs or HABs and then incubated at room temperature for 30 min. The samples were spun down at 100g for 30s, and resuspended with 30% (w/v) sucrose solution (Sigma, S0389) incubating overnight. After removing the supernatant, 200 µl OCT (Sakura, 4583) was added into samples. Samples were frozen in liquid nitrogen for 30 seconds. To obtain frozen sections, the frozen OCT block was transferred to a cryotome (Epredia™ Microm HM525 NX Cryostat, Germany) prior to sectioning and was allowed to equilibrate to same temperature as cryotome. Then, 5 ~ 10 µm thick sections were made, which were mounted onto adhesive microscope slides. The prepared slides can be stored at − 80°C for up to 12 months.

The frozen section slides were thawed at room temperature for 20 min. Slides were rehydrated in PBS for 10 minutes before immunofluorescence staining. A hydrophobic barrier was drawn around the sample slices' sections, followed by 100 ml of blocking buffer (5% BSA in PBST) and 30 min of incubation at room temperature. The primary antibody was then applied in the incubation buffer (1% BSA, 0.3% Triton X-100%, and 0.01% sodium aside in PBS) and incubated overnight at 4°C. The slides were then washed three times with PBS for 10 minutes each. The secondary antibody diluted in incubation buffer were applied and incubated at room temperature for 2h in darkness. Finally, the slides were washed by PBS for 10 min, and mounted with cover glass. The finished slides were analyzed with a confocal microscope (Nikon, Z2).

In situ immunofluorescent staining

HABs were fixed with 2.5% glutaraldehyde (Sigma-Aldrich, G5882) for 2 h and permeabilized with 0.1% Triton-X 100 (P0096, Beyotime) for 15 min. After washing with PBS, HABs were blocked with 10% (w/v) BSA (9048-46-8, Sigma-Aldrich) for 1 h. HABs were incubated with primary antibody for 12h at 4℃. After removing unconjugated primary antibody, samples were then incubated with the corresponding secondary antibodies and DAPI (Abcam, ab104139) for 2 h at room temperature while avoiding light. The control groups were obtained by replacing the primary antibodies with corresponding immunoglobulin G antibodies. Nikon Z2 confocal microscopy was used to capture all images. Primary antibodies and secondary antibodies used are listed in Supplementary Table S2.

Transport of fluorescent bile salt /CLF staining

HABs were incubated in Leibovitz L-15 medium (21083027, Gibco) containing 5µM cholyl-lysyl-fluorescein (CLF, BD-451041) for 15 min at 37℃ and 5% CO₂. Following two washes with L-15 medium, fluorescent bile salt internalization was visualized using a Nikon Z2 confocal microscope equipped with an incubator to maintain 37°C for observation.

Periodic Acid-Schiff Staining

PAS staining was used to study the accumulation of glycogen in liver. HABs were prepared and stained with PAS staining kits according to the manufacturer’s instructions (Solarbio, G1281). Glycogen accumulation in the scaffold was observed using a phase-contrast optical microscope (BX51, Olympus).

Transcriptome analysis

Novogene (China) performed RNA isolation, cDNA library construction, sequencing, and basic analysis. TRNZOL was used to collect samples for RNA-seq. According to the manufacturer's recommendations, 1µg of RNA per sample was employed to establish sequencing libraries using the NEBNext® UltraTM RNA Library Prep Kit for Illumina® (NEB, USA). Attribute sequences for each sample were assigned index codes. The Bioanalyzer 2100 system measured RNA integrity using the RNA Nano 6000 Assay Kit.

Sequencing was performed with an Illumina NovaSeq 6000 platform, and 150 bp paired-end reads were generated. Reads were filtered and mapped to the reference genome sequence with HISAT2 (version 2.0.5). FeatureCounts v1.5.0-p3 was used to count the read numbers mapped to each gene. The fragments per kilobase of exon per million fragments mapped (FPKM) method was used to calculate gene expression levels. Differentially expressed genes (DEGs) with |log2FoldChange|>0.0 and FDR q-value < 0.05 were identified using the DESeq2 R package (version 1.16.1), and the resulting p values were adjusted using Benjamini and Hochberg's approach for controlling the false discovery rate (FDR q-value). GO and KEGG enrichment analysis of DEGs were implemented by the cluster Profiler R package (version 3.8.1). Significantly enriched GO/KEGG terms were determined based on an adjusted p-value < 0.05. Significantly changed signaling pathways of GSEA were determined based on |NES|>1, FDR q-value < 0.25. Heatmaps were graphed by Heml 1.0.3.7-Heatmap illustrator. The PPI network was performed with Cytoscape ClueGO. Each term is represented by a circle node, whose size was proportional to the adjusted p-value.

Statistics

All data were presented as mean values ± standard deviation (SD). For comparisons between independent unpaired groups, an unpaired two-tailed student's t-test was performed when the variances were equal. An unpaired two-tailed Student's t-test with Welch's correction test was performed when the variances were unequal. For comparing multiple groups to a reference group, one-way ANOVA followed by Dunnett's test was used between groups with equal variance. After testing homogeneity of variance, LSD was used for pairwise comparison. Each experiment was performed in triplicate unless otherwise stated. Statistical significance was defined as * p < 0.05, ** p < 0.01, *** p < 0.001.

Data availability

The authors declare that all data supporting the findings of this study are available within the paper and its Supplementary Information or from the corresponding authors upon request.

Acknowledgment

The authors are sincerely grateful for funding from the National Key Research and Development Program of China (2022YFA1104600 and 2018YFA0109000) and the National Natural Science Foundation of China (31871015).

Author contribution

S.X. conducted all experiments and wrote the paper with input from all authors. Y.S. performed PMPs fabrication and characterization. C.X. assisted to perform the data visualization. R.Y. contributed to the study design and supervision. R.Y. and M.X. organized the project and provide financial support.

Competing interest

The authors declare no competing interests.

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Heterogeneous liver tissues with multicellular crosstalk and bile duct morphogenesis through organoid bioprinting

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Abstract

Figures

Introduction

Result And Discussion

The development of PMPs

Self-assembly and multicellular crosstalk of HOBBs

Reproducible bioprinting of HABs

Differentiation and liver function validation of HABs

HABs with bile duct morphogenesis

Conclusions

Materials And Methods

2D HepaRG cells culture

2D HUVECs culture and lentiviral transfection

PMP fabrication and mechanical testing

HOBB formation and culture

Determination of intracellular reactive oxygen species (ROS)

Live/Dead staining

Bioink preparation and characterization

3D bioprinting and culture of HABs

Flow Cytometry of Apoptosis of HABs

qRT-PCR Analysis

Cryosectioning & Immunofluorescent staining

Transport of fluorescent bile salt /CLF staining

Periodic Acid-Schiff Staining

Transcriptome analysis

Statistics

Declarations

References

Additional Declarations

Supplementary Files

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