Method for producing organoids using the self-organizing ability of stem cells and organoids produced by said method

Abstract

The present invention relates to a novel method for producing organoids using the self-assembly of spheroids. Specifically, the organoids produced by the method of the present invention have a cup-like shape with a concave center, which allows nutrients and oxygen to be smoothly supplied to the center, thereby overcoming the size limitations of organoids.

Description

[Technical Field]

[0001] The present invention relates to a novel method for producing an organoid tissue module using the self-assembly of spheroids, and to an organoid tissue module produced by said method. Specifically, the organoid tissue module produced by the production method of the present invention has a cup-like shape with a concave center, which allows nutrients and oxygen to be smoothly supplied to the center, thereby overcoming the size limitations of organoids. [Background technology]

[0002] Tissue engineering aims to restore or improve organ or tissue function by developing functional substitutes for damaged tissues (Griffith & Naughton, 2002; Langer & Vacanti, 1993). To develop such functional substitutes, research on organoids, which mimic human organs and tissues, is crucial. When stem cells are cultured in three-dimensional culture, they self-renew and self-organize to form organoids, three-dimensional cell aggregates composed of various cell types (Kaushik et al., 2018). Because the formed organoids can mimic the structural and functional properties of living tissues, they are useful not only for tissue regeneration therapy but also as miniaturized organ models for patient-customized disease modeling and drug screening systems (Kaushik et al., 2018).

[0003] Embryonic stem cells (ESCs), induced pluripotent stem cells (iPSCs), and adult stem cells (ASCs) are used as cell sources for organoids. Mesenchymal stem cells (MSCs), a type of adult stem cell, have attracted particular attention as a cell source for organoids because they can be easily isolated and cultured from various adult tissues, such as bone marrow and adipose tissue (Jiang et al., 2002; Pittenger et al., 1999; Wang et al., 2014). For 3D culture of these stem cells, a structural support similar to the extracellular matrix is ​​required. Matrigel is the most commonly used support, but its components are not fully characterized and it contains various growth factors that are difficult to quantify, making it difficult to maintain a consistent cell environment for each organoid culture. Alternatively, three-dimensional synthetic supports made of biomaterials such as hydrogels and polyethylene glycols can be used. However, these artificial supports have the disadvantage of not being able to fully reproduce the extracellular matrix of living tissues (Hofer & Lutolf, 2021; Khademhosseini & Langer, 2016). Therefore, there is an increasing demand for organoids to be generated without using artificial supports.

[0004] Embryonic tissue engineering overcomes this dependency on artificial supports by exploiting the ability of cells to synthesize their own extracellular matrix under specific conditions (Burdis & Kelly, 2021). When cells attach to the extracellular matrix they synthesize themselves, cell-cell and cell-extracellular matrix interactions occur, more similar to those found in real tissues. These interactions lead to biological processes such as self-assembly and self-organization. Self-assembly and self-organization processes can be induced using cell aggregation techniques. Most cell aggregation techniques rely on external physical forces, such as centrifugal force or electromagnetic fields, and most result in spherical structures. However, spherical organoids produced by these techniques are structurally limited in size due to the lack of sufficient oxygen and nutrients reaching the center. Therefore, long culture times are required to obtain large organoids, and increased culture times make it difficult to control the cellular composition of the organoids.

[0005] Therefore, in order to solve the above problems, the inventors developed a method for producing organoids that are not limited in size by utilizing the self-aggregation and self-organization capabilities of stem cells, and also isolated human mesenchymal stem cells (hMSCs) from human fat, then cultured them in two dimensions, and then used three-dimensional culture technology to create millimeter-sized organoid tissue modules made of mesenchymal stem cells, thereby completing the present invention.

[0006] As a result, the inventors produced millimeter-sized cup-shaped cartilage organoids in a chondrocyte differentiation environment and confirmed that transplanting the organoids into damaged articular cartilage resulted in cartilage regeneration. [Prior art documents] [Patent documents]

[0007] [Patent Document 1] Korean Patent Registration No. 10-2170076 Summary of the Invention [Problem to be solved by the invention]

[0008] The present invention provides a method for producing organoids of several millimeters in size by mixing and culturing spheroids of different sizes and utilizing the self-assembly ability of the spheroids. The method of the present invention simplifies the production method of complex organoids and provides organoids with a cup-like morphology that can supply nutrients and oxygen to the central region. [Means for solving the problem]

[0009] To achieve the above object, the present invention provides a method for producing an organoid tissue module by mixing and culturing spheroids of different diameters.

[0010] The present invention also provides an organoid tissue module produced by the above-described production method.

[0011] Furthermore, the present invention provides a pharmaceutical composition for preventing or treating cartilage diseases, comprising an organoid tissue module produced by the above-described production method.

[0012] In the above production method, the diameter of the spheroids is 10 to 1,000 μm, and preferably 50 to 500 μm.

[0013] In the above production method, spheroids having a diameter of 200 μm or less account for 60% or more of all spheroids, preferably 70% or more, and more preferably 80% or more.

[0014] In the production method, spheroids having a diameter of more than 200 μm account for 40% or less of all spheroids, preferably 30% or less, and more preferably 20% or less.

[0015] Furthermore, in the production method, the number of spheroids having a diameter of 200 μm or less may be greater than the number of spheroids having a diameter of more than 200 μm.

[0016] Furthermore, in the production method, the space occupied by spheroids having a diameter of 200 μm or less may be larger than the space occupied by spheroids having a diameter of more than 200 μm.

[0017] Furthermore, in the production method, the space not occupied by spheroids is 5 to 40% of the entire space in which the spheroids are cultured, preferably 10 to 35%, and more preferably 15 to 30%.

[0018] Furthermore, in the production method, the organoid tissue module may occupy 10 to 35% of the culture area, and the average diameter of the organoid tissue module may be 2 to 4 mm.

[0019] Furthermore, in the production method, the spheroids are obtained by culturing stem cells or undifferentiated cells, and the stem cells and undifferentiated cells are preferably selected from the group consisting of embryonic stem cells, induced pluripotent stem cells, adult stem cells, and mesenchymal stem cells, and more preferably, the stem cells are human adipose-derived mesenchymal stem cells (hMSCs).

[0020] Furthermore, in the manufacturing method, the organoid tissue module may be cup-shaped.

[0021] Additionally, the manufacturing method may be free of scaffolds and artificial materials. [Effects of the Invention]

[0022] The present invention provides a technique for producing organoid tissue modules and tissue composites that are not restricted by scaffolds and xenotransplantation. The organoid tissue modules produced by the production method of the present invention are cup-shaped, eliminating the problem of oxygen and nutrient supply. Organoid tissue modules can be produced easily and in a short time by utilizing the self-assembly ability of spheroids.

[0023] Furthermore, organoids produced by the production method of the present invention maintain the properties of stem cells and can therefore be used to treat a variety of diseases, including cartilage disorders. [Brief explanation of the drawings]

[0024] [Figure 1] FIG. 1 is a schematic diagram showing the principle of the technology for producing organoid tissue modules (Tissue modules; TM). [Figure 2] 1 shows photographs and graphs showing the morphology of human mesenchymal stem cells (hMSCs) and their cell division ability through subculture. [Figure 3] 1 is a graph showing positive stem cell surface markers of hMSCs using FACS. [Figure 4] 1 is a graph showing stem cell surface negative markers of hMSCs using FACS. [Figure 5] This is a photograph showing microscopic confirmation that hMSCs have differentiation potential properties. [Figure 6] This is a photograph of a spheroid mixture that failed to generate an organoid tissue module. [Figure 7] This is a photograph of a spheroid mixture that successfully generated an organoid tissue module. [Figure 8] These are photographs and schematic diagrams of time-lapse images taken at various times during the formation of organoid tissue modules over a 72-hour period. [Figure 9]This photograph shows the process of organoid tissue module formation, in which smaller spheroids are attracted to and fused with larger spheroids at the center. [Figure 10] 1 is a graph showing the size distribution of spheroids produced for the production of organoid tissue modules. [Figure 11] In order to successfully produce organoid tissue modules, the spheroid mixtures that successfully produced organoid tissue modules were classified into sizes with a diameter of 200 μm or less and sizes with a diameter of more than 200 μm, and the graph shows the average diameter of spheroids in each range. [Figure 12] 1 is a graph showing the distribution of spheroid numbers at each size for mixed spheroids produced using the organoid tissue module and mixed spheroids not produced using the organoid tissue module. [Figure 13] 10 is a graph showing the spatial distribution of spheroids of each size for mixed spheroids produced using the organoid tissue module and mixed spheroids not produced using the organoid tissue module. [Figure 14] 1 is a graph showing the distribution of unoccupied space in mixed spheroids produced using the organoid tissue module and in mixed spheroids not produced using the organoid tissue module. [Figure 15] 1 is a graph showing the average movement speed of spheroids over time during the process of organoid tissue module formation by self-organization. [Figure 16] 1 is a graph showing the deviation of the migration distance and average speed of spheroids over time during the process of forming an organoid tissue module by self-organization. [Figure 17] 1 is a graph showing the trajectory of spheroids during the process of forming organoid tissue modules through self-organization. [Figure 18]1 is a graph showing the circularity of organoid tissue module formation stages, the rate at each time, and the time to reach the initiation, condensation, lifting, and folding stages. [Figure 19] 1 shows photographs and graphs showing that adjusting the cell number changes the size of spheroids. [Figure 20] 10 shows photographs showing the effect of the mixing ratio of two types of spheroids with different sizes on the formation of organoid tissue modules. [Figure 21] 10 shows photographs showing whether or not organoid tissue modules were generated at various ratios of the area not occupied by mixed spheroids to the total culture area. [Figure 22] These photographs show that organoid tissue modules made from two types of spheroids of different sizes self-organize and maintain their cup-shaped morphology. [Figure 23] Photographs showing the process of fusion of two types of spheroids of different sizes. [Figure 24] This is a photograph of a fluorescently stained organoid tissue module taken with a confocal microscope. [Figure 25] 1 shows photographs and graphs showing the results of examining the stem cell markers and differentiation potential of cells isolated from organoid tissue modules. [Figure 26] Photographs and graphs showing that organoid tissue modules are resistant to cell death in comparison with spherical cell aggregates. [Figure 27] 1 shows photographs confirming the presence or absence and location of type II collagen expression by immunohistological staining in an organoid tissue module in which cartilage differentiation was induced. [Figure 28] 1 shows photographs and graphs showing aggrecan expression and the distribution of nuclei stained with SYTO16 after 3D clearing of an organoid tissue module in which chondrogenic differentiation was induced. [Figure 29]1 is a graph showing the glycosaminoglycan (GAG) content of the cartilage extracellular matrix in an organoid tissue module in which chondrogenic differentiation was induced. [Figure 30] Photographs showing the results of histological analysis of organoid tissue modules in which cartilage differentiation was induced, using various staining methods. [Figure 31] Photographs showing the efficacy and histological analysis results of transplantation of an organoid tissue module in which cartilage differentiation has been induced into rabbit cartilage. DETAILED DESCRIPTION OF THE INVENTION

[0025] DETAILED DESCRIPTION OF THE PREFERRED EMBODIMENTS

[0023] The present invention will now be described in detail with reference to the accompanying drawings, so that those skilled in the art can easily understand the present invention. However, the present invention may be embodied in various forms and is not limited to the embodiments and examples set forth herein.

[0026] Throughout this specification, when a part is said to "comprise" a certain element, this does not mean that it excludes other elements, but that it may further include other elements, unless otherwise specified.

[0027] The organoid tissue module manufacturing method of the present invention includes a first step of isolating mesenchymal stem cells (hMSCs) from human adipose tissue, followed by two-dimensional culture and three-dimensional culture techniques to produce spheroids; and a second step of mixing and culturing spheroids of various sizes to produce an organoid tissue module of cup-shaped organoids.

[0028] A schematic diagram of the present invention for generating organoids of several millimeters in size from single cells is shown in Figure 1.

[0029] The term "two-dimensional culture" refers to culturing cells in a monolayer in various types of culture vessels, such as cell culture flasks or flat Petri dishes.

[0030] The term "three-dimensional culture" refers to culturing cells while allowing them to interact with all three dimensions surrounding them, and is characterized by the ability of cells to grow in all directions.

[0031] The term "organoid" refers to a tissue mimic that is a three-dimensional culture of stem cells and exhibits a morphology similar to that of actual human tissues and organs through self-organization.

[0032] The "organoid tissue module (Tissue Module;™)" refers to an organoid that has not differentiated into a specific organ or tissue and is produced by self-organization of spheroids.

[0033] The term "cup shape" means a shape having a recess in the middle.

[0034] The "stem cells" of the present invention include adult stem cells, and are preferably "mesenchymal stem cells", and more preferably "human adipose-derived mesenchymal stem cells".

[0035] The present invention will be described in more detail below with reference to examples. However, these examples are merely for illustrating the present invention and are not intended to limit the present invention. [Example]

[0036] Isolation of mesenchymal stem cells (hMSCs) from adipose tissue Human adipose-derived mesenchymal stem cells (hMSCs) were used as the cell source for the production of the organoid tissue module of the present invention, as they are relatively easy to isolate and have a high growth rate (Meligy et al., 2012). Isolation of hMSCs was performed with the approval of the Institutional Review Board of Seoul National University Dental Hospital.

[0037] The prepared human adipose tissue was placed in Hanks' Balanced Salt solution (HBSS) containing 0.1% collagenase I and shaken at 37°C for 30 minutes to separate the cells. The cells were then washed and filtered through a 100 μm nylon mesh. The cells were then supplemented with Dulbecco's modified Eagle's medium (DMEM) containing 10% fetal bovine serum (FBS) and 1× antibiotic-antimycotic solution (AA) and cultured at 37°C in a 5% CO2 incubator. [Example]

[0038] Validation of isolated human mesenchymal stem cells (hMSCs) As mentioned above, isolated hMSCs have characteristics such as adherent morphology, multipotency, and expression of surface-specific antigens (Dominici et al., 2006). Therefore, by confirming these characteristics, we verified that the cells isolated in Example 1 were hMSCs.

[0039] The morphology and distribution characteristics of the cells were confirmed, and as shown in Figure 2, the cells adhered to and grew in a uniform spindle-like shape on a plastic culture dish, and the time it took for the number of cells to double was consistently 2 days from the second to the 12th passage.

[0040] To confirm whether the characteristics of hMSCs were well maintained after repeated passages, the expression of stem cell surface markers was examined by fluorescence microscopy. At passages 3, 5, and 7, cells were incubated with fluorescently labeled antibodies against CD14, CD29, CD31, CD34, CD44, CD73, CD90, HLA-DR, CD45, CD79α, CD105, and CD117 for 1 hour at 4°C. After washing with Dulbecco's phosphate buffered saline (DPBS) containing 2% FBS, the expression levels were analyzed by FACS compared with an isotype mouse IgG control.

[0041] As a result of this test, as shown in Figure 3, more than 95% of the cells expressed positive markers CD29, CD44, CD73, CD90, and CD105, and as shown in Figure 4, it was confirmed that the expression of negative markers CD14, CD31, CD45, CD79α, CD117, and HLA-DR was less than 2%. [Example]

[0042] Confirmation of the osteocyte differentiation potential of isolated mesenchymal stem cells (hMSCs) To confirm the osteogenic differentiation potential of hMSCs, 9.5 × 10 4 hMSCs were cultured in alpha minimum essential medium (αMEM) containing 10 nM dexamethasone, 50 μg / mL ascorbic acid, 10 mM β-glycerolphosphate, 1 mM dibutyryl-cAMP, 10% FBS, and 1x AA. After 4 days, the culture medium was removed and replaced with dibutyryl-cAMP-free medium twice a week. The cells were cultured at 37°C in a 5% CO2 incubator for 14 days to induce osteogenic differentiation.

[0043] To visually confirm osteogenic differentiation, alizarin red S staining was performed. After differentiation, the culture medium was removed from the plates, and the plates were washed once with DPBS. Then, 1 mL of 4% paraformaldehyde (PFA) was added and the cells were fixed at room temperature. The fixative was removed, and the plates were washed with DPBS. 2 mL of alizarin red S solution diluted to 1% with distilled water (DW) was added to each plate and incubated for 15 minutes at room temperature. The plates were then washed five times with DW, and 1 mL of DW was added. Osteogenic differentiation was confirmed under a microscope (Olympus) as shown in Figure 5. [Example]

[0044] Confirmation of the adipocyte differentiation potential of isolated human mesenchymal stem cells (hMSCs) To confirm the adipocyte differentiation ability, 9.5 × 10 cells were cultured in the same manner as in Example 3. 4 Cells were plated in 6-well plates and cultured in a culture medium containing 0.5 mM 3-isobutyl-1-methylxanthine (IBMX), 100 nM dexamethasone, 100 μM indomethacin (INDO), 10 μg / mL insulin, 10% FBS, and 1x AA in high-glucose DMEM for 14 days in a 5% CO2 incubator at 37°C with medium changes twice a week. To confirm differentiation, the culture medium was removed from the plates after differentiation, washed once with DPBS, and then fixed with 4% PFA at room temperature.

[0045] To visually confirm differentiation, 0.2 g of oil red O powder was dissolved in 40 mL of isopropanol and filtered through a 0.22 μm filter to prepare a solution (1 mL). The cells were then incubated at room temperature for 15 minutes, washed five times with DW, and the cells were then confirmed to have differentiated into adipocytes (adipogenic differentiation) using an Olympus microscope, as shown in Figure 5. [Example]

[0046] Confirmation of the chondrocyte differentiation potential of isolated human mesenchymal stem cells (hMSCs) To confirm the chondrocyte differentiation ability, 9.5 × 10 4The cells were placed in a 6-well plate and cultured in a culture medium prepared by adding 1x insulin-transferrin-selenium premix (ITS), 50 ng / mL ascorbic acid, 40 μg / mL L-proline, 100 nM dexamethasone, 10% FBS, and 1x AA to α-MEM, supplemented with 10 ng / mL transforming growth factor (TGF-β3). The cells were then cultured at 37°C in a 5% CO2 incubator for 21 days. After differentiation, the culture medium was removed from the plate, and the cells were fixed with 4% PFA at room temperature.

[0047] After fixation, the cells were washed with DPBS, placed in OCT compound, and frozen in a deep freezer (-80°C) for 1 day. The frozen cells were then cut into 8 μm-thick sections using a cryocut microtome and washed with DW. A 1% alcian blue solution prepared using alcian blue reagent was added to the sections, stained at room temperature for 30 minutes, and then washed with 0.1 N HCl solution and DW. Chondrogenic differentiation was confirmed under an Olympus microscope, as shown in Figure 5.

[0048] The differentiation ability was confirmed in Examples 3 to 5, and it was confirmed that the cells isolated from human fat using the technology of the present invention were of extremely high purity and retained the characteristics of stem cells even after long-term culture. [Example]

[0049] Preliminary research for the creation of organoid tissue modules Generally, organoid generation requires long-term culture, and as organoids grow larger, the supply of oxygen and nutrients to cells located in the center of the spherical organoid becomes limited, increasing the likelihood of cell death. Therefore, we sought to identify conditions for generating cup-shaped organoid tissue modules that would ensure a smooth supply of oxygen and nutrients to the center. After identifying the conditions for generating organoid tissue modules, we then attempted to provide a differentiation environment during the generation process of the organoid tissue modules, thereby rapidly generating organoids that grew to several millimeters in size.

[0050] Using the hMSCs prepared and confirmed in the above examples, spheroids of various sizes were generated for the construction of organoid tissue modules. 10,000 or 15,000 hMSCs were placed in each well of a 96-well flat or U-bottom plate, and forced aggregation was induced by centrifugation at 500 x g for 5 minutes. The cells were then cultured to produce randomly aggregated cell aggregates, which were then cultured to produce spheroids of various sizes.

[0051] Because hMSC spheroids ranging in size from 50 to 500 μm in diameter formed in each well, a 96-well plate was used to contain all of the spheroids. After centrifugation, the supernatant was removed and the spheroids were resuspended in 200 μl of medium and transferred to a single well of an ultra-low attachment 96-well plate. The spheroids were then placed in a 37°C incubator to induce self-assembly.

[0052] The size range of the generated spheroids varied depending on the condition of the bottom of the 96-well plate used (flat or round bottom). As shown in Figure 6, there were wells where no self-assembly occurred, while there were wells that contained organoid tissue modules with a unique structure that self-assembled and had a central depression, as shown in Figure 7. These cup-shaped structures with a unique self-assembly form, measuring 2-4 mm in diameter, were named organoid tissue modules. [Example]

[0053] Analysis of spheroid organoid tissue module generated images The spheroids were mixed and transferred to a plate of designated size (e.g., a 96-well plate). Time-lapse images were taken from the time the spheroids sank to the bottom of the well (time 0) for 72 hours. Each time-lapse image was combined to animate the spheroid self-assembly process, which was then analyzed. As shown in Figure 8, the spheroids actively moved and passed through the initiation stage, condensation stage, and lifting and folding stage, resulting in the formation of a cup-shaped organoid tissue module.

[0054] The time when the spheroids sank to the bottom of the well (time 0) was designated the initiation phase. The stage in which each spheroid moved to the center and aggregated was designated the condensation phase. The stage in which each spheroid formed a cup-like shape was designated the lifting and folding phase. In the initiation phase, each spheroid had a clearly separated boundary. In the condensation phase, spheroids moved to the center, fused with neighboring spheroids, and continued to shrink toward the core. In the lifting and folding phase, the edges of the organoid tissue module lifted and bent upward against gravity, generating a cup-shaped organoid tissue module.

[0055] As shown in Figure 9, when large and small spheroids aggregated, the smaller spheroids were attracted to the larger spheroids and aggregated. These results indicate that the size of the spheroids and the proportion of each size are important in inducing self-organization for organoid tissue modules. [Example]

[0056] Spheroid size analysis for organoid tissue module production To confirm the spheroid mixing conditions for generating organoid tissue modules, we analyzed images of the initiation stage in wells that successfully and unsuccessfully generated organoid tissue modules.

[0057] The diameter of the measurable spheroids at the initiation stage was determined using Image J, and the spheroids were classified into size ranges as shown in Table 1 and Figure 10.

[0058] [Table 1]

[0059] The average diameter of spheroids in each size range was 80 μm for spheroids under 100 μm, 140 μm for spheroids between 100 and 200 μm, 250 μm for spheroids between 200 and 300 μm, 330 μm for spheroids between 300 and 400 μm, and 430 μm for spheroids between 400 and 500 μm.

[0060] When the size of the spheroids in the wells in which the organoid tissue modules were generated was analyzed, as shown in Figure 11, the average diameter of small spheroids with a diameter of 200 μm or less was 110 μm, and the average diameter of large spheroids with a diameter of more than 200 μm was 270 μm. [Example]

[0061] Analysis of the number distribution of spheroids in each size range for organoid tissue module production The results of analyzing the number of spheroids of each size that constituted wells in which organoid tissue modules were successfully generated (Success) are shown in Table 2 and Figure 12.

[0062] [Table 2]

[0063] The results of analyzing the number of spheroids of various sizes that constituted wells in which organoid tissue modules were not produced (Failure) are shown in Table 3 and Figure 12.

[0064] [Table 3]

[0065] As shown in Tables 2 and 3, when the spheroid diameters mixed at the organoid tissue module initiation stage were various sizes ranging from 50 to 500 μm, it was confirmed that organoid tissue modules were not produced when 23% of the spheroids were 200 μm or less, but were produced when 82% of the spheroids were 200 μm or less.

[0066] It was also confirmed that organoid tissue modules were not produced when 77% of the spheroids had a diameter of more than 200 μm, but were produced when 18% of the spheroids had a diameter of more than 200 μm.

[0067] Furthermore, it was confirmed that organoid tissue modules were produced when the number of spheroids with a diameter of 200 μm or less was greater than the number of spheroids with a diameter of more than 200 μm. [Example]

[0068] Spatial occupancy analysis of spheroids for organoid tissue module generation To confirm the conditions for creating organoid tissue modules, the occupied space and residual space of spheroids of each size were analyzed.

[0069] The results of analyzing the occupied space of each size of spheroids constituting wells in which organoid tissue modules were successfully generated (Success) are shown in Table 4 and Figure 13. The occupied space is expressed as a percentage of the total space occupied by spheroids belonging to each size range relative to the total space occupied by all spheroids.

[0070] The space occupied by each spheroid was calculated using the radius (r) obtained from Image J analysis using the formula π × r 2 was calculated using

[0071] [Table 4]

[0072] Table 5 and Figure 13 show the results of analyzing the space occupied by spheroids of each size that constituted wells in which organoid tissue modules were not produced (Failure).

[0073] [Table 5]

[0074] As shown in Tables 4 and 5, when the spheroid diameters mixed at the initiation stage of the organoid tissue module were various sizes ranging from 50 to 500 μm, it was confirmed that an organoid tissue module was not produced when spheroids of 200 μm or less occupied 6% of the total spheroid space, but was produced when they occupied 48%.

[0075] It was also confirmed that organoid tissue modules were not produced when spheroids with a diameter of more than 200 μm occupied 94% of the total spheroid space, but were produced when they occupied 52%.

[0076] The remaining space, excluding the space occupied by the spheroids, was further analyzed using Image J software, assuming the bottom area of ​​the well to be 100%. The results are shown in Figure 14 and Table 6.

[0077] As shown in Figure 14, Tables 6 and 7, when organoid tissue modules were produced, it was confirmed that the area of ​​empty space relative to the total area was 5 to 40%.

[0078] In other words, it can be seen that the area of ​​free space required for producing an organoid tissue module is 5 to 40%, preferably 10 to 35%, and more preferably 15 to 30%.

[0079] [Table 6]

[0080] [Table 7] [Example]

[0081] Bio-kinematic analysis of organoid tissue module construction To confirm the self-assembly process of the organoid tissue module, biokinematic analysis was performed as follows.

[0082] The movement of spheroids during the organoid tissue module formation process was videotaped, and the spatiotemporal distribution of spheroids' speed, trajectory, linearity, and coordination was analyzed and calculated based on the video. From the images taken over time, the motility of spheroids within a limited space during the formation of the organoid tissue module was calculated using the Image J program and a MATLAB code based on the particle image velocimetry (PIV) algorithm.

[0083] As shown in Figure 15, typical spheroid movement characteristics were observed during the initiation, condensation, lifting, and folding stages of organoid tissue module generation. The fluctuations in the average spheroid velocity every 12 hours were due to the culture medium changes performed every 12 hours, and the changes in motility due to the medium changes were excluded from the analysis.

[0084] In the initial stage, each spheroid exhibited irregularly varying motility speeds (0-1.5 μm / min).

[0085] During the condensation stage, the movement of the central part of the organoid tissue module was relatively static, while the spheroids in the boundary region still exhibited dynamic motility.

[0086] During the lifting and folding stages, most spheroid movements were reduced and stabilized, with some repetitive movements observed only in the boundary region.

[0087] Furthermore, analysis of the spheroid migration distance (path length) showed that the boundary regions of the organoid tissue module during the condensation stage exhibited linear movement, whereas the spheroids in the central region exhibited twisted movement, as shown in Figure 16. During the lifting and folding stages, spheroid movement was barely observed in the boundary regions.

[0088] Trajectory analysis (Figure 17) showed that spheroids exhibited random movement during the initiation stage, whereas spheroids in the boundary region were observed to move linearly toward the center during the condensation stage. Uniquely, spheroids in the folding region exhibited longer trajectories and greater linearity than those in the center. This type of movement is generally observed when aggregates form stable structures, indicating that the organoid tissue module in the lifting and folding stages was transformed into a structurally stable structure.

[0089] Furthermore, we analyzed phenotypic parameters such as circularity. As shown in Figure 18, when the spheroids were mixed and cultured for 24 hours, the circularity of the organoid tissue module temporarily decreased due to spheroid motility. However, after an additional 18 hours of culture, the circularity approached 1.0, which means a circle with the same diameter in all directions. These results indicate that the organoid tissue module, which has an irregular structure, begins to reorganize into a regular structure within 24 hours.

[0090] Furthermore, as shown in Figure 18, biokinetic and morphological analyses reveal that the organoid tissue module self-organizes through stages of initiation (0 h), condensation (4.3 h), lifting and folding (39 h). [Example]

[0091] Organoid Tissue Module Creation Using Optimized Conditions The experimental results of Examples 8 to 11 confirmed that the mixing ratio of spheroids with a diameter of 200 μm or less to spheroids with a diameter of more than 200 μm, the spatial occupancy rate of spheroids with a diameter of 200 μm or less to spheroids with a diameter of more than 200 μm, and the area of ​​free space relative to the space in which the organoid tissue module is cultured are directly related to the generation of the organoid tissue module.

[0092] Therefore, in order to confirm the above conditions, experiments were carried out under various conditions.

[0093] Each microwell of the AggreWell plate was filled with 100, 500, 1000, 3000, or 5000 cells, and the culture medium was DMEM containing 40 μg / mL L-proline, 100 μg / mL sodium pyruvate, 50 μg / mL L-ascorbic acid 2-phosphate, 1×ITS, and 1×AA.

[0094] The results of culturing in Aggrewell are shown in Figure 19. For spheroids with an average diameter of 200 μm or less, spheroids consisting of 500 cells were selected. For spheroids with an average diameter of more than 200 μm, spheroids consisting of 3,000 cells were selected. Spheroids consisting of 500 cells were named 0.5K, and spheroids consisting of 3,000 cells were named 3K spheroids.

[0095] As shown in Example 10, the two sizes of spheroids obtained as described above, 0.5K and 3K, were mixed and arranged in varying proportions in an ultra-low attachment 96-well plate so that the free area was approximately 20% of the total culture area.

[0096] First, when the concentration of 0.5K spheroids was increased to 10%, 30%, or 50%, self-assembly did not occur. In contrast, when 0.5K and 3K spheroids were mixed at 60% or more of 0.5K spheroids and 40% or less of 3K spheroids, organoid tissue modules were formed through self-assembly of the spheroids, as shown in Figure 20. Therefore, when spheroids of different diameters are mixed and cultured, organoid tissue modules are produced, and when the number of spheroids with a diameter of 200 μm or less is greater than the number of spheroids with a diameter of more than 200 μm, organoid tissue modules are successfully produced.

[0097] To investigate the effect of free space on organoid tissue module formation, the ratio of 0.5K spheroids to 3K spheroids was fixed at 90% and 10%, respectively, and the free culture area not occupied by spheroids was adjusted to 90%, 80%, 70%, 60%, 50%, 40%, 30%, 20%, 10%, and less than 10%, as shown in Figure 21. It was confirmed that when the free area was greater than 50%, spheroid self-assembly was insufficient, while when the free area was less than 40%, organoid tissue modules were successfully formed by spheroid self-assembly.

[0098] In another example, the area of ​​free space was set to 20% of the total culture area, and then the spheroids were cultured to induce self-organization to produce organoid tissue modules. As shown in Figure 22, the organoid tissue modules were generated during the self-assembly process, and it was confirmed that they maintained their cup-like shape even after 7 days of culture.

[0099] Furthermore, during the process of organoid tissue module generation, spheroids fused with neighboring spheroids and condensed to form larger structures. When spheroids fused to form organoid tissue modules, the cells constituting the spheroids were not uniformly mixed and remained as the original spheroids, as confirmed by staining the spheroid fusion process, as shown in Figure 23. After surface fluorescent staining, confocal microscopy confirmed that the organoid tissue modules were cup-shaped, as shown in Figure 24. The average diameter of the organoid tissue modules was approximately 2-4 mm (10-35% of the culture area), approximately half the size of the pineal gland (5-8 mm), the smallest organ in the human body. [Example]

[0100] Confirmation of hMSC stem cell properties of the created organoid tissue module To confirm whether the hMSCs constituting the organoid tissue module maintained their stem cell properties, single cells were isolated from the organoid tissue module and examined for stem cell marker expression and differentiation potential. As shown in Figure 25, the cells constituting the organoid tissue module expressed stem cell markers, differentiated into adipocytes, osteocytes, and chondrocytes, and maintained their stem cell properties. [Example]

[0101] Confirmation of cell death resistance of the created organoid tissue module To confirm whether the organoid tissue modules prepared were more resistant to cell death than spherical cell aggregates, we prepared spherical aggregates and organoid tissue modules with the same number of cells. For the spherical cell aggregates, cells were placed in a U-bottom plate and centrifuged at 500 × g for 5 minutes. Then, cells were induced to aggregate in an incubator at 37°C and cultured for a total of 6 days under the same conditions as the organoid tissue modules. While the spherical cell aggregates exhibited significant cell death and surrounding dead cell debris, the organoid tissue modules exhibited relatively minimal cell debris, as shown in Figure 26. The organoid tissue modules and spherical cell aggregates were fixed in 4% PFA for 1 day and embedded in paraffin blocks. Slides were prepared at 4 μm thickness, deparaffinized, and then treated with proteinase K to dissolve the tissue proteins. Endogenous peroxidase was removed by treatment with 3% hydrogen peroxide (H2O2), and TUNEL (Terminal Deoxynucleotidyl Transferase dUTP Nick-end Labeling) staining was performed using the ApoTag Peroxidase In Situ Apoptosis Detection Kit (Millipore). Nuclei were counterstained with methyl green, and images were then acquired. As shown in Figure 26, no increased cell death was observed in the center of the organoid tissue module compared to the periphery. However, there were many dead TUNEL-stained cells in the center of the spherical cell aggregates, and empty spaces were formed due to necrosis. Therefore, the organoid tissue module's cup-shaped structure, with its concave center, appears to facilitate the smooth supply of nutrients and oxygen, providing a technical advantage that overcomes the disadvantage of increased central cell death with increasing size. [Example]

[0102] Confirmation of the possibility of differentiation of the created organoid tissue module into chondrocytes by checking the presence or absence of Type 2 collagen expression. The hMSCs used in this study are stem cells that differentiate into chondrocytes and have been used as a promising cell source for cartilage tissue engineering (Somoza et al., 2014). To confirm the potential of the organoid tissue modules prepared as described above to differentiate into chondrocytes, the organoid tissue modules were treated with TGF-β3 to induce chondrocyte differentiation. The samples were then collected and fixed in 4% PFA for approximately 3 days, followed by immunohistochemistry (IHC) for histological analysis.

[0103] To prepare slides, fixed samples were embedded in paraffin blocks and 4 μm thick slides were prepared. The slides were deparaffinized in histo-clear for 30 minutes, hydrated in 100% ethanol and then 70% ethanol, and then washed with PBS. Blocking was then performed for 1 hour at room temperature with 0.3% bovine serum albumin (BSA) in PBS. An appropriate concentration of primary antibody against type 2 collagen was then diluted in the blocking solution and allowed to stand at 4°C for at least 12 hours. After the blocking period, the slides were washed with PBS, and an appropriate concentration of secondary antibody was added to the blocking solution. The slides were then allowed to stand at 37°C for 1 hour. After washing with PBS, the slides were then reacted with DAB for color development.

[0104] The results of this experiment are shown in Figure 27. It was confirmed that type 2 collagen is mainly expressed in the spherical cells that correspond to large spheroids. [Example]

[0105] Confirmation of the potential for differentiation of the created organoid tissue module into chondrocytes by Aggrecan confirmation To confirm aggrecan, a marker of chondrocyte differentiation, the spheroids were cleared by a 3D clearing process according to the method described in Patent Document 1 (Method for producing decellularized tissue using hydrogel polymer and decellularized tissue produced thereby), and then the expression of aggrecan was examined in 3D using an antibody.

[0106] To achieve this, the organoid tissue modules were fixed in 4% PFA for one week, washed with 0.1% Triton-X in PBS, and then blocked for at least 8 hours at room temperature using a solution of 6% BSA, 0.2% Triton X-100, and 0.01% sodium azide in PBS. Aggrecan antibodies were then diluted to the appropriate concentration in the blocking solution, dispensed into the organoid tissue modules, and incubated at room temperature for two days. After washing with the aforementioned washing solution, secondary antibodies and Hoechest were diluted to the appropriate concentrations in the blocking solution, dispensed into the spheroids, and incubated at room temperature for two days. After the antibody reaction, the spheroids were cleared using a solution of 25% urea and 65% sucrose in water.

[0107] The results of this experiment are shown in Figure 28. Aggrecan expression was confirmed to be highest at the edges of the organoid tissue module. [Example]

[0108] Quantitative comparative analysis of glycosaminoglycan-induced cartilage differentiation Glycosaminoglycans (GAGs), an important extracellular matrix component of cartilage, were used in a quantitative comparative analysis of cartilage differentiation.

[0109] To prepare a papain solution, 7.8 μL of papain was added to a solution of 200 mM sodium phosphate, 100 mM sodium acetate, 10 mM EDTA, and 5 mM L-cysteine, adjusted to pH 6.4. The samples were then incubated in a constant-temperature water bath at 65°C for 18 hours. After 18 hours, the mixture was centrifuged at 10,000 g for 10 minutes, and the GAG ​​content was measured according to the sulfate glycosaminoglycan assay kit guidelines.

[0110] The absorbance of the samples was measured at 656 nm using a microplate reader, and the total DNA amount was quantified using a pico-green dsDNA assay kit for normalization of the measurement results.

[0111] The results of the experiment are shown in Figure 29. It was confirmed that the GAG ​​content was significantly increased in the organoid tissue module compared to single cells. [Example]

[0112] Histological analysis of organoid tissue modules induced to differentiate into cartilage using various staining methods To confirm the immunohistochemical staining results, organoid tissue module samples were embedded in paraffin blocks and sliced ​​into 4 μm-thick slides. After deparaffinization as described above, the slides were stained with hematoxylin and eosin (H&E). Synthesized collagen in the organoid tissue modules was detected using the Trichrome III Blue Staining Kit, and GAG expression was visualized using the Safranin-O / fast green kit.

[0113] The results of this experiment are shown in Figure 30. Masson's trichrome staining revealed blue staining throughout the organoid tissue module due to collagen synthesis, while Safranin-O staining confirmed that red-stained GAGs were expressed throughout the organoid tissue module. This confirms that the organoid tissue module differentiates into chondrocyte organoids in the presence of chondrogenic differentiation factors such as TGF-β3. [Example]

[0114] Efficacy of organoid tissue module confirmed using a rabbit cartilage defect model To confirm the cartilage regeneration efficacy of organoid tissue modules in a cartilage defect osteoarthritis (CD / OA) model, we created a cartilage defect model by removing the entire cartilage layer from the knee of adult New Zealand White rabbits and transplanted organoid tissue modules into the defect site. This experiment was conducted under the approval of the Seoul National University Hospital Institutional Animal Care and Use Committee (SNUH-IACUC No. 18-0171-S1A0), and animals were maintained in a facility assessed and certified by the Association for Assessment and Accreditation of Laboratory Animal Care (AAALAC) in accordance with the National Research Corporation (NRC) Laboratory Animal Care and Use Guide, 8th Edition.

[0115] Rabbits were anesthetized by intramuscular injection of a mixture of 5 mg / kg xylazine and 15 mg / kg zoletil, and a 3 mm diameter x 3 mm deep cartilage defect was induced in the knee joint using sterile surgical tools. The organoid tissue module transplantation group received the organoid tissue module, which was then sutured. For 3 days after surgery, the animals were treated with antibiotics and analgesics via subcutaneous injection of 0.3 mL Metacam and 1 mL Cefazolin. The animals were sacrificed 12 or 14 weeks after surgery, and the knees were resected and subjected to micro-computed tomography (micro-CT) imaging prior to histological evaluation.

[0116] After the experiment, the rabbit cartilage was photographed using Micro-CT, and it was confirmed that all of the defect areas had been completely filled, as shown in Figure 31. In addition, paraffin sections were prepared using rabbit knee cartilage tissue and H&E staining was performed. This confirmed that the formation of epiphyseal plate structures had increased in the defected cartilage, and the formation of extensive extracellular matrix and bone lacunae was clearly observed at the transplant site of the organoid tissue module.

[0117] Furthermore, the site was confirmed by safranin-O staining, and as a result, as shown in Figure 31, the formation of hyaluronic cartilage was confirmed in the organoid tissue module transplant group.

[0118] The experiment showed that the cartilage generated 12 weeks after transplantation of the organoid tissue module into the CD / OA rabbit model was thicker than the surrounding cartilage, and that the organoid tissue module had completely integrated with the damaged cartilage. This indicates that the millimeter-sized organoid tissue module has an excellent regenerative effect in the CD / OA model.

Claims

1. A method for producing organoid tissue modules by mixing and culturing spheroids of different diameters.

2. A method for producing an organoid tissue module according to claim 1, characterized in that the diameter of the spheroids is 10 to 1,000 μm.

3. A method for producing an organoid tissue module as described in claim 1, characterized in that the diameter of the spheroids is 50 to 500 μm.

4. A method for producing an organoid tissue module as described in claim 1, characterized in that spheroids with a diameter of 200 μm or less account for more than 60% of all spheroids.

5. A method for producing an organoid tissue module as described in claim 1, characterized in that spheroids with a diameter of 200 μm or less account for more than 70% of all spheroids.

6. A method for producing an organoid tissue module as described in claim 1, characterized in that spheroids with a diameter of 200 μm or less account for more than 80% of all spheroids.

7. A method for producing an organoid tissue module as described in claim 1, characterized in that spheroids with a diameter of more than 200 μm account for 40% or less of the total spheroids.

8. A method for producing an organoid tissue module as described in claim 1, characterized in that spheroids with a diameter of more than 200 μm account for 30% or less of the total spheroids.

9. A method for producing an organoid tissue module as described in claim 1, characterized in that spheroids with a diameter of more than 200 μm account for 20% or less of the total spheroids.

10. A method for producing an organoid tissue module as described in claim 1, characterized in that the number of spheroids with a diameter of 200 μm or less is greater than the number of spheroids with a diameter of more than 200 μm.

11. A method for producing an organoid tissue module as described in claim 1, characterized in that the space not occupied by spheroids is 5 to 40% of the total space in which the spheroids are cultured.

12. A method for producing an organoid tissue module as described in claim 1, characterized in that the space not occupied by spheroids is 10 to 35% of the total space in which the spheroids are cultured.

13. A method for producing an organoid tissue module as described in claim 1, characterized in that the space not occupied by spheroids is 15 to 30% of the total space in which the spheroids are cultured.

14. A method for producing an organoid tissue module as described in claim 1, characterized in that the tissue module occupies 10 to 35% of the culture area.

15. A method for producing an organoid tissue module as described in claim 1, characterized in that the diameter of the tissue module is approximately 2 to 4 mm.

16. A method for producing an organoid tissue module as described in claim 1, characterized in that the spheroids are obtained by culturing stem cells or undifferentiated cells.

17. A method for producing an organoid tissue module as described in claim 16, characterized in that the stem cells or undifferentiated cells are selected from the group consisting of embryonic stem cells, induced pluripotent stem cells, adult stem cells, and mesenchymal stem cells.

18. A method for producing an organoid tissue module as described in claim 17, characterized in that the stem cells are human adipose-derived mesenchymal stem cells (hMSCs).

19. A method for manufacturing an organoid tissue module as described in claim 1, characterized in that the organoid tissue module is cup-shaped.

20. A method for producing an organoid tissue module as described in claim 1, characterized in that the method does not include scaffolds and artificial substances.

21. An organoid tissue module produced by the production method described in any one of claims 1 to 20.

22. A pharmaceutical composition for preventing or treating a cartilage disease, comprising the organoid tissue module described in claim 21.

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