Organoids can be used for emulating the complex process of tissue and organ development in vitro. Stem cells in three-dimensional (3D) culture can give rise to self-organizing multicellular structures termed organoids that can resemble the anatomical and functional units of the organ from which they are derived. As organoids can recapitulate the complexity of in vivo physiological systems at the convenience of in vitro cell culture, they can be used for modeling healthy and/or diseased states of various adult organs for biomedical and pharmaceutical applications.
To provide a 3D environment for the organoid culture, certain techniques require embedding stem cells in sessile drops of extracellular matrix (ECM) hydrogels prepared from solubilized basement membrane extracts (e.g., Matrigel). When supplied with culture media containing defined soluble factors that permit proper cell growth and directed differentiation into organ-specific lineages, the 3D environment can induce the differentiating cells to segregate into distinct domains and undergo fate specification, leading to their spontaneous organization into organ-like structures. Despite its utility and versatility, however, these techniques can be limited due to the limited lifespan of organoids. In a typical setup, developing organoids embedded in an ECM hydrogel rely on passive diffusion for nutrient supply and waste removal. This mode of transport effectively supported the organoids through the hydrogel scaffold at their initial stages of development. As the organoids grow and become more metabolically demanding, however, limited diffusion of nutrients and oxygen into the inner regions of the 3D scaffold causes a progressive and significant reduction in the viability of organoids, resulting in the formation of a necrotic core. The rate at which this degenerative process occurs varies depending on the type of organoids, but in most cases, considerable cell death becomes evident within 10 days of culture using the ECM hydrogel. Certain culture techniques can avoid cell death by passaging organoids every 5-7 days. However, such a short duration of each cycle can interrupt the continuous culture of organoids for prolonged periods necessary for their sustained development and maturation into in vivo-like tissue constructs.
To address this problem, researchers have used bioreactors to improve diffusive transport of oxygen and nutrients in 3D culture of organoids. As demonstrated by recent work on cerebral organoids, this approach has proven instrumental for establishing long-term culture to promote continued development and increased maturity of organoids. Implementing this technique in routine laboratory settings, however, is burdened by the need for capital equipment that is mechanically complex and requires specialized knowledge for operation and maintenance. Another drawback is that organoids are cultured in suspension in bioreactors, which makes it challenging to monitor their growth and development during culture. While vascularization of organoids has been suggested as an alternative strategy to improve nutrient supply, the process of generating organoid models with controlled vascular perfusion is prohibitively complex and often requires advanced in vitro systems and specialized techniques that are not easily accessible to non-engineers.
Therefore, there is a need for improved techniques that can be used for uninterrupted and continuous culture of organoids for prolonged periods.
The disclosed subject matter provides techniques for culturing organoids and/or cells. An example device for culturing organoids can include an access port configured to receive a solution, a loading chamber, and a plurality of culture chambers. In non-limiting embodiments, the access port can be located in the center of the loading chamber. In some embodiments, the culture chambers can be radiated from the loading chamber so that the solution injected into the loading chamber through the access port can be evenly distributed into the culture chambers. In non-limiting embodiments, the culture chambers can be open to an external environment and include a protruding edge at an opening of the culture chambers.
In certain embodiments, the device can include poly(dimethylsiloxane). In non-limiting embodiments, the device can be optically transparent.
In certain embodiments, the solution can be a hydrogel solution. In non-limiting embodiments, the hydrogel solution can include cells and/or organoids. In some embodiments, the organoid can be a human organoid. In non-limiting embodiments, each culture chamber can include a different type of cells or organoids for co-culturing. In some embodiments, at least about 80% of the organoids in the culture chamber can be viable at day 21 of culturing. In non-limiting embodiments, the growth of the organoids can continue for at least 21 days. In some embodiments, the size of the organoids can increase for at least 21 days. In certain embodiments, the device can reduce the variability in the size of the organoids.
In certain embodiments, each of the culture chambers can have a width and a height ranging from about 100 μm to about 5 cm. In non-limiting embodiments, the protruding edge can be configured to pin a meniscus of the solution at the opening of the culture chambers for filing the entire culture chambers without spillage of the solution through the open-top.
The disclosed subject matter also provides methods for culturing organoids. An example method can include injecting a hydrogel precursor solution including organoids into a loading chamber through an access port, filling a plurality of culture chambers with the hydrogel precursor solution including organoids, solidifying the hydrogel precursor solution to form a hydrogel in the plurality of culture chambers, and providing culture media that is contacted to the hydrogel through the open-top. In non-limiting embodiments, the access port can be located in the center of the loading chamber. In some embodiments, the culture chambers can be radiated from the loading chamber so that the hydrogel precursor solution injected into the loading chamber can be evenly distributed into the culture chambers. In non-limiting embodiments, the culture chambers can be open to an external environment and include a protruding edge at an opening of the culture chambers for preventing spillage of the hydrogel precursor solution through the open-top.
In certain embodiments, the culture media includes soluble factors. In non-limiting embodiments, the soluble factors can include a growth factor, an active agent, or a combination thereof.
In certain embodiments, the method can further include maturing the organoids. In non-limiting embodiments, the method can further include assessing the viability and maturation of the organoids in the plurality of the culture chamber. The plurality culture chamber can be transparent. In some embodiments, the organoid can be a human organoid.
According to an embodiment, the present disclosure relates to a device for culturing organoids, comprising an access port configured to receive a solution, a loading chamber, wherein the access port is located in the loading chamber, and a plurality of culture chambers, wherein the culture chambers are radiated from the loading chamber so that the solution injected into the loading chamber through the access port is distributed into the plurality of culture chambers, wherein the plurality of culture chambers are open to an external environment and comprises a protruding edge at an opening of the plurality of culture chambers.
In an embodiment, the device comprises poly(dimethylsiloxane). In an embodiment, the device is optically transparent. In an embodiment, the access port is located in a center of the loading chamber. In an embodiment, the plurality of culture chambers are symmetrical with respect to rotations about the access port. In an embodiment, the solution injected into the loading chamber through the access port is evenly distributed into the plurality of culture chambers. In an embodiment, the device is configured to contact a culture media from the external environment through the opening of the plurality of culture chambers. In an embodiment, the solution is a hydrogel solution. In an embodiment, the hydrogel solution comprises cells or organoids. In an embodiment, the organoids are human organoids. In an embodiment, each of the culture chambers has a width or a height ranging from about 100 μm to about 5 cm. In an embodiment, each of the culture chambers has a width and a height of about 1 cm. In an embodiment, at least about 80% of the organoids in the culture chamber are viable at day 21 of culturing. In an embodiment, the protruding edge is configured to pin a meniscus of the solution at the opening of the culture chambers, allowing filling of the culture chambers without spillage of the solution through the opening. In an embodiment, each culture chamber comprises a different type of cells or organoids for co-culturing. In an embodiment, growth of the organoids continues for at least about 21 days. In an embodiment, a size of the organoids increases for at least about 21 days. In an embodiment, the device decreases variability in the size of the organoids.
According to an embodiment, the present disclosure relates to a method for culturing organoids, comprising injecting a solution including cells or organoids into a loading chamber through an access port, filling a plurality of culture chambers with the solution including cells or organoids, wherein the culture chambers are radiated from the loading chamber so that the solution injected into the loading chamber is distributed into the plurality of culture chambers, wherein the plurality of culture chambers are open to an external environment and comprises a protruding edge at an opening of the culture chambers for preventing spillage of the solution through the opening, and providing a culture media to the device through the opening of the plurality of culture chambers.
In an embodiment, the culture media comprises soluble factors. In an embodiment, the soluble factors are selected from the group consisting of a growth factor, an active agent, and a combination thereof. In an embodiment, the method further comprises maturing the organoids. In an embodiment, the method further comprises assessing viability and maturation of the organoids in the plurality of culture chambers.
The disclosed subject matter will be further described below.
It is to be understood that both the foregoing general description and the following detailed description are exemplary and are intended to provide further explanation of the disclosed subject matter.
The disclosed subject matter provides techniques for culturing cells and/or organoids. The disclosed techniques can provide enhanced organogenesis and extended life span of the cells or organoids. The disclosed techniques can also enhance the maturity of the cells and organoids. The disclosed techniques can also permit the enlargement of the cells and organoids. The disclosed techniques can also reduce the variability of the cells and organoids.
Unless otherwise defined, all technical and scientific terms used herein have the same meaning as commonly understood by one of ordinary skill in the art. In case of conflict, the present document, including definitions, will control. Certain methods and materials are described below, although methods and materials similar or equivalent to those described herein can be used in the practice or testing of the presently disclosed subject matter. All publications, patent applications, patents, and other references mentioned herein are incorporated by reference in their entirety. The materials, methods, and examples disclosed herein are illustrative only and not intended to be limiting.
As used herein, the term “organoid” generally describes a 3D multicellular in vitro tissue construct that mimics its corresponding in vivo organ such that it can be used to study aspects of that organ. As used herein, the term “organoid” describes any geometry of self-organized three-dimensional tissue culture. In certain instances, the term “organoid” may be further defined as comprising stem cells and/or somatic cells.
As used herein, the term “about” or “approximately” means within an acceptable error range for the particular value as determined by one of ordinary skill in the art, which will depend in part on how the value is measured or determined, i.e., the limitations of the measurement system. For example, “about” can mean within 3 or more than 3 standard deviations, per the practice in the art. Alternatively, “about” can mean a range of up to 20%, up to 10%, up to 5%, and up to 1% of a given value. Alternatively, particularly with respect to biological systems or processes, the term can mean within an order of magnitude, within 5-fold, and within 2-fold, of a value.
The present disclosure introduces a facile, scalable engineering approach to enable long-term development and maturation of organoids. Method described herein have redesigned the three-dimensional configuration of conventional organoid culture to develop a platform that converts single injections of stem cell suspensions to radial arrays of organoids that can be maintained for extended periods. Using human and mouse stem cells, accelerated production of intestinal organoids and their sustained development for over 4 weeks without the need for passaging is demonstrated. Compared to conventional techniques, long-term culture in the disclosed device enhances the formation of the crypt-villus structures and significantly increases the functional maturity of the intestinal epithelium. Further, vascularized, perfusable human enteroids can be assembled in a microengineered device and used to model the recruitment of innate immune cells to the diseased intestinal epithelium in IBD. The disclosed system, methods, and device may provide an immediately deployable platform to engineer more realistic organ-like structures in a dish.
The present disclosure describes a simple, immediately deployable strategy based on rethinking the design of conventional 3D culture of organoids. The methods described herein utilize an advanced platform capable of reconfiguring the geometry of 3D culture scaffolds to generate open arrays of organoids that eliminate the problem of limited and non-uniform diffusion inherent in bulk hydrogel. The systems described herein can be manufactured as simple and ready-to-use culture inserts of different sizes and shapes that can be used in standard cell culture plates without any modification of established protocols and workflow.
A proof-of-concept of the present disclosure is demonstrated by continuous growth and development of mouse intestinal organoids during extended periods of uninterrupted culture. Resulting intestinal tissue constructs exhibit structural and functional maturity not achievable in conventional culture. The utility of the approach can be further demonstrated by the production and prolonged maintenance of human intestinal organoids and in-depth analysis of their significantly enhanced maturity using single-cell RNA sequencing (scRNA-seq). Finally, the advanced capabilities of the technology can be demonstrated by establishing i) an organotypic model of human inflammatory bowel disease (IBD) using patient-derived enteroids and ii) microengineered organoids integrated with perfusable vasculature for in vitro modeling of immune-epithelial interaction in IBD.
As background, it should be appreciated that central to the challenge of long-term culture in traditional organoid models is the formation of areas of increased cell death due to diffusion limitations in the inner regions of sessile hydrogel drops. In essence, the approach described herein can be conceptualized as i) removing this necrotic core from the hydrogel scaffold while keeping the outer layer in which organoids remain viable and ii) radially segmenting this layer and spreading it out to form a planar array of organoids, as shown in
For practical implementation of this idea, a disc-shaped 3D culture device that enables the production and maintenance of radially arranged organoid arrays in standard cell culture plates was created (
In addition to procedural simplicity and convenience, OCTOPUS enables new capabilities that render the method advantageous over conventional techniques. The design of OCTOPUS as a removable culture insert makes this system easily transferable (
Briefly introduced above, the system, methods, and device of the present disclosure will now be described in more detail below with reference to the Drawings.
In certain embodiments, the disclosed subject matter provides a device for culturing cells, organoids, or tissue explants. As shown in
In certain embodiments, the loading chamber 102 can be configured to receive a solution through the access port 101. For example, a solution can be pipetted into the loading chamber 102 through the access port 101. In non-limiting embodiments, the access port 101 can be located in the center of the loading chamber 102. In some embodiments, the device can include more than one loading chamber 102 for co-culturing different types of cells and organoids. In certain embodiments, the loading chamber 102 can include poly(dimethylsiloxane) (PDMS). In certain embodiments, the loading chamber 102 can include polystyrene, thermoplastic, glass, metal, paper, or combinations thereof.
In certain embodiments, the loading chamber 102 can have a diameter ranging from about 2 mm to about 10 mm. In non-limiting embodiments, the access port 101 can have a diameter ranging from about 0.5 mm to about 3 mm.
In certain embodiments, the culture chamber 103 can be radiated from the loading chamber 102 so that the solution injected into the loading chamber 102 through the access port 101 can be evenly distributed into the at least one culture chamber 103. In non-limiting embodiments, multiple culture chambers 103 can be radiated from the loading chamber 102. In some embodiments, the disclosed device can include more than one loading chamber 102, and each loading chamber can be connected to one or more culture chambers for the co-culturing platform. Each culture chamber 103 can include a different type of cells or organoids, while they can be exposed to the same culture media. Each culture chamber 103 can include a different type of extracellular matrix. In certain embodiments, each culture chamber 103 can contain independently accessible flow channels. In certain embodiments, the culture chamber 103 can include PDMS. In certain embodiments, the culture chamber 103 can include polystyrene. In certain embodiments, the culture chamber 103 can include thermoplastics. In certain embodiments, the culture chamber 103 can include glass. In certain embodiments, the culture chamber 103 can include metals. In certain embodiments, the culture chamber 103 can include paper.
In some embodiments, the culture chamber 103 can have a width ranging from about 100 μm to about 50 mm. In non-limiting embodiments, the culture chamber 103 can have a height ranging from about 100 μm to about 5 cm. In some embodiments, the shape and size of the culture chamber 103 or can be modified depending on the purposes of the disclosed device (e.g., co-culture, target cells, and target organoids). In some embodiments, the culture chamber 103 can have a width and/or height of about 100 μm to about 5,000,000 μm. In some embodiments, the culture chamber 103 can have a width and/or height of at least about 100 μm. In some embodiments, the culture chamber 103 can have a width and/or height of at most about 5,000,000 μm. In some embodiments, the culture chamber 103 can have a width and/or height of about 100 μm to about 1,000 μm, about 100 μm to about 10,000 μm, about 100 μm to about 50,000 μm, about 100 μm to about 100,000 μm, about 100 μm to about 500,000 μm, about 100 μm to about 1,000,000 μm, about 100 μm to about 5,000,000 μm, about 1,000 μm to about 10,000 μm, about 1,000 μm to about 50,000 μm, about 1,000 μm to about 100,000 μm, about 1,000 μm to about 500,000 μm, about 1,000 μm to about 1,000,000 μm, about 1,000 μm to about 5,000,000 μm, about 10,000 μm to about 50,000 μm, about 10,000 μm to about 100,000 μm, about 10,000 μm to about 500,000 μm, about 10,000 μm to about 1,000,000 μm, about 10,000 μm to about 5,000,000 μm, about 50,000 μm to about 100,000 μm, about 50,000 μm to about 500,000 μm, about 50,000 μm to about 1,000,000 μm, about 50,000 μm to about 5,000,000 μm, about 100,000 μm to about 500,000 μm, about 100,000 μm to about 1,000,000 μm, about 100,000 μm to about 5,000,000 μm, about 500,000 μm to about 1,000,000 μm, about 500,000 μm to about 5,000,000 μm, or about 1,000,000 μm to about 5,000,000 μm. In some embodiments, the culture chamber 103 can have a width and/or height of about 100 μm, about 1,000 μm, about 10,000 μm, about 50,000 μm, about 100,000 μm, about 500,000 μm, about 1,000,000 μm, or about 5,000,000 μm.
In certain embodiments, the culture chambers can be open to an external environment. As shown in
In certain embodiments, the culture media can include nutrients, soluble factors, growth factors, active agents, or combinations thereof. For example, when supplied with culture media containing the soluble/growth factors that permit proper cell growth and directed differentiation into organ-specific lineages, the cells and/or organoids in the 3D culture chamber can be differentiated into organ-like structures. In certain embodiments, the culture media can include soluble/growth factors such as R-spondin ligand, Noggin, bone morphogenetic protein (BMP), epithelial growth factor (EGF), fibroblast growth factor (FGF), B-27, N-2, BSA, ascorbic acid, MTG, Glutamax, CHIR99021, rhKGF, 8BrcAMP, IBMX, DMH-1, A83-01, hydrocortisone, and heparin. In non-limiting embodiments, the culture media can include a target active agent for screening drugs. For example, intestinal stem cells can be seeded into the culture chamber 103 and treated with anti-fibrotic drugs (e.g., Pirfenidone and/or Nintedanib) at pre-determined concentrations for testing the effects of the drugs on the fibrotic phenotype. In non-limiting embodiments, the active agent can include chemicals, toxins, nanomaterials, bacteria, viruses, nucleic acids, peptides, or combinations thereof.
In non-limiting embodiments, the culture chamber 103 can include a protruding edge 106, or step 106, at the opening 104 of the culture chamber 103. The protruding edge 106 can be configured to pin a meniscus of the injected solution at the opening-top of the culture chamber 103 for filing the entire culture chamber 103 without spillage of the solution through the open-top.
In certain embodiments, the culture chamber 103 can be coated for enhancing the adhesion of a gel and/or a cell to the inner surface of the culture chamber 103. For example, each culture chamber 103 can be filled with a dopamine hydrochloride solution at room temperature (RT) to form a surface coating for enhanced adhesion of a hydrogel.
In certain embodiments, the disclosed device can include PDMS. In certain embodiments, the loading chamber can include polystyrene. In non-limiting embodiments, the device can be optically transparent. For example, cells or organoids embedded in a hydrogel located in the disclosed device can be observed through microscopic techniques (e.g., bright-field, confocal, fluorescence, electron, atomic force, and laser scanning microscopy) without removing the hydrogel from the disclosed device. In certain embodiments, the device can have a size ranging from about 1 mm to about 50 cm.
In certain embodiments, the solution injected into the loading chamber can be a hydrogel solution. For example, the hydrogel solution can be an extracellular matrix (ECM) precursor solution, which can be solidified (i.e., gelation) after in the culture chamber, providing 3D culture environments. In non-limiting embodiments, the solution can include cells, organoids, or tissue explants. The cells can be any cells that can be cultured in vitro. For example and not limitation, the cells can be stem cells, goblet cells, endothelial cells, epithelial cells, mesenchymal cells, neural cells, muscle cells, progenitor cells, immune cells, endocrine cells, or combinations thereof. The organoids can be any organoids that can be cultured in vitro. For example and not limitation, the organoids can include human organoids, mouse organoids, intestinal organoids, liver organoids, lung organoids, nascent organoids, or combinations thereof.
In certain embodiments, the organoids cultured in the disclosed device can have an extended life span. For example, The organoids cultured in the disclosed system can survive up to about 3 weeks without passaging. In non-limiting embodiments, at least about 80% of the organoids in the culture chamber can be viable at day 5, 10, 14, and 21 of culture.
In certain embodiments, the disclosed device can provide improved morphological and functional maturation of the organoids. The long-term culture capabilities of the disclosed device can be leveraged to increase the maturity of intestinal organoids. For example, during culture (e.g., for about 7 days), the flat epithelium can be folded into finger-like protrusions (e.g., villi) and have extended budding, which can be longer than the villi cultured without the disclosed device. In addition to morphological development, the disclosed device can provide improved functional maturation of organoids. For example, the villi cultured in the disclosed device can express higher functional markers (e.g., peptide transporter 1, sodium-glucose linked transporter 1 (SGLT1), and glucose transporter 2 (GLUT2)) than the villi cultured without the disclosed device.
In certain embodiments, the disclosed subject matter provides methods for culturing organoids. An example method can include injecting a hydrogel precursor solution including organoids into a loading chamber through an access port, filling a plurality of culture chambers with the hydrogel precursor solution including organoids, solidifying the hydrogel precursor solution to form a hydrogel in the plurality of culture chambers, and providing culture media that is contacted to the hydrogel through the open-top. For example, to form organoids in the disclosed device, the organoid/hydrogel mixture can be generated by mixing the hydrogel precursor solution with a pellet of organoids in a complete organoid growth medium. Using a pre-wetted pipet tip, about 100 μl of the organoid/Matrigel mixture can be injected into the disclosed device through the access port. The mixture can be evenly distributed through the culture chamber without spillage of the mixture solution. For example, each culture chamber can have the same volume of the mixture after being injected through the access port. The disclosed device can be incubated for gelation of the hydrogel precursor solution. Pre-warmed organoid growth media can be added to each culture chamber for long term culture.
In certain embodiments, the method can further include assessing the viability and maturation of the organoids in the plurality of the culture chamber through the transparent device. For example, viability and maturation of the organoids can be assessed through microscopic techniques (e.g., bright-field, confocal, fluorescence, electron, atomic force, and laser scanning microscopy) and biochemical analyses (e.g., ELISA).
According to an embodiment, a device according to the above can be fabricated by casting PDMS prepolymer against micropatterned three-dimensional printed molds using standard soft lithography techniques. For example, PDMS (Sylgard 184, Dow Corning, USA) monomer base can be mixed with a curing agent (10:1, w/w) and poured onto 3D printed molds (Protolabs, USA). The casted molds can be vacuum degassed in a desiccation chamber for 30 minutes, after which the PDMS can be oven cured overnight at 65° C. to produce devices containing organoid culture chambers, as described in
The disclosed device can extend the lifespan of organoids. To simulate the most common settings of standard organoid culture, small intestinal organoids derived from commercially available mouse adult stem cells and protocols were selected as a model system.
In an attempt to provide further insight into the observed differences, temporal profiles of 70-kDa FITC-dextran permeation into organoid-containing Matrigel scaffolds to simulate passive diffusion of soluble factors in the disclosed device was measured. For spatial analysis, these measurements were taken at two locations representing the inner and outer regions of the scaffold.
These results suggest that the 3D culture environment in the device can allow for unrestricted and spatially uniform diffusive transport of soluble factors. In Matrigel sessile drops, unrestricted transport of dextran into the outer layer was evident from the rapid increase in fluorescence intensity within 30 minutes, which was in contrast to limited dye penetration into the core of the scaffold (
Consistent with these findings, oxygen penetration into the Matrigel scaffold in OCTOPUS occurred in a rapid and uniform manner, resulting in oxygen saturation through the entire thickness of the construct within 30 minutes (
These results verify the design principle of OCTOPUS and suggest that long-term viability and sustained growth of organoids with reduced size variability shown in the disclosed system may be attributed to unrestricted and spatially uniform diffusive transport of nutrients, growth factors, and oxygen, which is achieved by decreasing the distance between organoids and the Matrigel surface (
Limited lifespan of organoids in conventional culture hampers their ability to reach later stages of development and acquire a more mature phenotype. The long-term of OCTOPUS can be leveraged to increase the maturity of intestinal organoids in the model system.
During embryogenesis, the flat epithelium of the developing gut tube begins to fold into finger-like protrusions called the villi, which are separated by deep invaginations known as the crypts (
Results also showed higher expression of stem cell markers (Lgr5, Ki67) (
In addition to morphological development, how the 3D culture environment in OCTOPUS affects the emergence of organ-specific tissues in intestinal organoids was assessed. Stem and progenitor cells in the embryonic intestine give rise to a specialized epithelium on the villi that contain absorptive and secretory cells critical for nutrient absorption and other important physiological functions of the intestine. Indeed, this process of epithelial differentiation and maturation occurred in both conventional and OCTOPUS organoid models as evidenced by the expression of hepatocyte nuclear factor 4α (Hnf4α), which is a transcription factor that plays an essential role in intestinal maturation during embryogenesis. Organoids in Matrigel drops, however, were seen with considerably lower expression even at the point of their maximum maturation (day 7) (
Importantly, the enhanced maturation of organoids in the device was further supported by analysis of cell type-specific markers. At day 7, for example, the expression of villin—a terminal differentiation marker of absorptive enterocytes—was significantly upregulated in OCTOPUS compared to hydrogel drops (
In the next phase, whether the enhanced morphogenesis and tissue maturity demonstrated in the device contribute to the functional maturation of intestinal organoids was assessed. Given that the primary function of the intestine is nutrient absorption, key molecular transporters that regulate the absorptive function of the intestinal epithelium, including i) peptide transporter 1 (PEPT1) responsible for intestinal uptake of peptides and ii) sugar transporters including sodium-glucose linked transporter 1 (SGLT1) and glucose transporter 2 (GLUT2) that mediate the absorption of monosaccharides were measured.
In this analysis, organoids in hydrogel drops at the maximum duration of culture (7 days) were compared to those maintained in OCTOPUS for 14 days to examine the contribution of extended culture. Regardless of the culture platform, immunostaining clearly showed the presence of the transporters on the villi, but the expression of these functional markers was significantly elevated in OCTOPUS (
For further functional characterization, live-cell imaging techniques were used to visualize intracellular calcium signaling, which has been shown to regulate the activity of the intestinal nutrient transporters detected in the model. To assess the level of intracellular calcium, the organoids were labeled with a fluorescent calcium indicator dye (Fluo-4) and monitored their fluorescence in real-time. Given the multicellular complexity of organoids, the mean fluorescence intensity was measured from representative organoids selected for analysis. Upon treatment with 100 μM ATP, organoids in the device increased their fluorescence by approximately 1.6-fold within 60 seconds, which was followed by a gradual decrease to the baseline level (
The results have shown that intracellular calcium signaling also plays an essential role in the secretion of digestive hormones in response to increased nutrients in the intestinal lumen, which is another important physiological function of the intestine. Based on this evidence, hormone secretion in the disclosed organoid models was assessed as a measure of functional maturation. Enzyme-linked immunosorbent assay (ELISA) of conditioned media was performed to measure the glucose-induced release of glucagon-like peptide 1 (GLP-1), an incretin hormone secreted by enteroendocrine L cells of the intestinal epithelium that enhances glucose-stimulated insulin release from pancreatic β-cells. The ELISA data showed the release of the biologically active form of GLP-1 by the cultured intestinal organoids in response to glucose included in the culture media. Notably, for all three-time points of analysis (days 5, 7, and 10), the hormone was secreted in significantly larger amounts in OCTOPUS than was measured in the conventional model (
While organoids have the inherent capacity to reproduce the multicellular complexity of their in vivo counterparts, it remains a significant challenge to emulate the integrated higher-level structure and function of native organs in conventional organoid culture. To meet this challenge, efforts are being made to develop new methods for increasing the cellular heterogeneity of current organoid models and recapitulating biological crosstalk beyond the cellular level of organization to model tissue-tissue and multiorgan interactions. Inspired by this emerging body of work, the possibility of using OCTOPUS to create co-culture models that combine organoids with their associated tissues in 3D culture was assessed.
First, the design of OCTOPUS was engineered to incorporate a pair of open spiral culture chambers with individually accessible injection ports (
The dual-chamber design could easily be modified during device fabrication to accommodate a greater number of tissue types. This was demonstrated by increasing the number of chambers to create a tri-culture system that consisted of intestinal organoids and two neighboring 3D constructs containing intestinal fibroblasts and blood vessels (
Importantly, analysis of the co-culture models indicated significant effects of non-parenchymal tissues on the development of organoids. For example, when a co-culture of small intestinal organoids with primary intestinal fibroblasts was established (
Recognizing that in vitro modeling of complex disease is emerging as the primary focus of organoid research, the potential application of OCTOPUS for this active area of investigation was assessed. With the goal of exploiting the long-term culture capabilities of the disclosed system, a model intestinal fibrosis was established as a representative example of pathophysiological conditions caused by prolonged disease processes that cannot be easily recapitulated in conventional organoid models due to their limited lifespan. Fibrosis is a common complication of intestinal diseases such as inflammatory bowel disease and gastrointestinal cancer. The intrinsic ability of the intestine to repair wounds and restore homeostasis can be impaired by repetitive epithelial injury due to persistent insults such as chronic inflammation. The dysregulated process of wound healing can lead to abnormal remodeling of the sub-epithelial tissue characterized by the activation of fibroblasts and excessive deposition of ECM. One of the goals was to construct an organoid-based advanced in vitro model capable of emulating these salient features of fibrotic tissue remodeling in the intestine.
To this end, OCTOPUS was used to set up a co-culture of intestinal organoids and primary intestinal fibroblasts in the same hydrogel scaffold and generate a multicellular construct reminiscent of the intestinal epithelium and its underlying stroma in vivo (
To simulate the scenario of using this system to assess intestinal fibrosis in a conventional laboratory setting, a common technique widely used for in vitro modeling of fibrosis, which was to treat the co-culture construct with transforming growth factor (TGF)-β, were used. TGF-β plays a central role in the pathogenesis of fibrosis in the intestine and other organs by inducing the activation of fibroblasts and their transdifferentiation into myofibroblasts, which are the key effector cells that drive fibrogenesis. To trigger fibrogenic responses, the model was treated with TGF-β at 1 ng/ml from day 5 to day 12. Exposure of the intestinal microtissue to this condition indeed caused the fibroblasts to acquire the contractile phenotype of myofibroblasts, as evidenced by robust expression of alpha-smooth muscle actin (αSMA) (
The model also permitted the investigation of ECM deposition, which is essential to fibrotic tissue remodeling. This analysis focused on fibronectin (FN) as a representative ECM protein.
During the same treatment period (from day 5 to day 12), immunostaining showed that stimulation with TGF-β substantially increased FN in the pericellular regions of fibroblasts as compared to the untreated tissues (
To further validate the fibrotic phenotype of the model, the stiffness of the TGF-β-treated microtissues was measured by using atomic force microscopy (AFM). This measurement was greatly facilitated by the open-top design of OCTOPUS that allowed direct access of the AFM probe to the tissue constructs in the culture chambers (
The utility of the intestinal fibrosis model for drug testing applications was tested. Given that no specific therapies are currently available for intestinal fibrosis, two anti-fibrotic drugs, Pirfenidone and Nintedanib, approved for the treatment of idiopathic pulmonary fibrosis (IPF), were used. Although these drugs were developed for fibrosis in the lung, they can modulate the activity of fibrogenic pathways in other organs such as the heart, kidney, liver, and skin. These findings led to an examination of whether the com-pounds would have similar therapeutic effects on intestinal fibrosis. The potential of the drugs to reverse the fibrotic phenotype of the model, namely fibroblast activation and excessive ECM deposition, was assessed.
First, fibrotic intestinal tissue constructs in OCTOPUS were generated by forming co-culture organoids over 5 days and exposing them to TGF-β for another 7 days as described above (day 5-day 12). These constructs were then treated with clinically relevant concentrations of the drugs for 48 hours (day 13-day 14) within the therapeutic window identified by viability assessment (data not shown). In a control group, the fibrotic tissues did not receive drug treatment during the 48-hour period. At 0.1 mM, Pirfenidone was effective for altering the contractile phenotype of fibroblasts, as illustrated by 50% reduction in αSMA compared to the untreated control (
In the case of Nintedanib, a lower dose of the drug (0.1 μM) failed to exert significant effects on the fibrosis model (
Consistent with the results of immunofluorescence analysis, AFM data demonstrated the anti-fibrotic effects of Pirfenidone and Nintedanib in a dose-dependent manner (
In response to the increasing need for new technologies for organoid research, here a microengineered platform was established to reconfigure the three-dimensionality of conventional organoid culture. OCTOPUS introduced in this paper provides a simple yet effective means to address the problem of limited nutrient supply inherent in 3D culture and engineer a more uniform, unrestricted soluble environment beneficial for long-term culture of organoids. As demonstrated by the model systems, the extended lifespan of organoids significantly increased their size and maturity beyond what is achievable using conventional techniques and enabled the production of more realistic multicellular constructs for in vitro modeling of organogenesis and disease development.
Organoids in conventional hydrogel drop scaffolds can be passaged weekly for prolonged periods to increase their in vitro lifespan. Mechanically disrupted organoids during subculture have the capacity to rapidly seal themselves and restore their original architecture and functional properties. As demonstrated by the long-term culture of intestinal organoids for over 1 year, this approach has proven instrumental for expanding organoids and maintaining their differentiated phenotype over extended periods. The increased lifespan of organoids in this case, however, does not necessarily translate into enhanced tissue maturity because frequent passaging (typically every 5-7 days) required by conventional culture protocols disrupts the process of sustained organoid development and maturation. OCTOPUS resolves this issue by enabling uninterrupted, continuous culture of organoids for significantly longer (>3×) periods of time.
Although the ability to support continuous long-term culture is the key advantage of OCTOPUS, the data also revealed other desirable features of organoid development in this system. After 7 days of culture, for example, virtually every marker of structural and functional maturation measured was expressed in significantly higher levels in OCTOPUS-generated or-ganoids (
By leveraging these capabilities, the work also demonstrated the feasibility of developing a specialized organoid model in OCTOPUS that can simulate the salient features of dysregulated fibrogenesis during the development of intestinal fibrosis. The disclosed subject matter entailed a sequential process of generating co-culture organoid constructs and then exposing them to a fibrogenic factor, which took place over time periods (12-14 days) well beyond the typical lifespan of organoids in conventional culture. Indeed, a rapid loss of organoid viability in Matrigel drops after day 5 made it challenging, if not impossible, to model fibrogenic responses to TGF-β (data not shown), which began to occur only after 8 days of continuous culture in OCTOPUS and became more evident over time (
While OCTOPUS represents considerable changes to the design of traditional organoid models, the implementation of this system does not require any modification of established culture protocols and workflow, nor does it rely on specialized equipment or personnel. Essential to this advantage is the design of OCTOPUS as a ready-to-use and easily-accessible culture insert that is directly compatible with standard well plates and laboratory infrastructure. As exemplified by the intestinal models, generating mature organoids in OCTOPUS can readily be accomplished in traditional laboratory settings based on materials and experimental procedures commonly used in conventional techniques. This is an important aspect of the method that makes OCTOPUS an immediately deployable and readily accessible culture platform, which can contribute to the rapid dissemination of the technology for widespread use.
The foregoing demonstration raises several fundamental questions that open new avenues for further investigation. Among them is which design parameters play a significant role in the long-term development of organoids in OCTOPUS. During extended culture, intestinal organoids in the device continued to grow in size as shown in
Perhaps the more important question is how long the system can support continuous growth and maturation of organoids before passaging becomes necessary. Many intestinal organoids in the device ceased to grow after 4 weeks of culture (data not are shown), which can be considered conservatively as the maximum duration of continuous culture of small intestinal organoids in the current design of OCTOPUS. This result begs the question of whether the arrested growth of organoids is due to the physical constraints of the culture chambers described above. Another explanation is that the dead cells sloughing off of the epithelium during the natural process of epithelial turnover and accumulating in the closed liminal cavity can exert deleterious effects on the organoids, which has been described previously in the same type of organoids. It can be that as organoids grow beyond a certain limit, the system in its current configuration reaches its maximum capacity and becomes no longer capable of meeting the metabolic demands of enlarged organoids. As discussed above, analysis of organoid development in culture chambers with different sizes and geometry can help address some of these questions. The external environment of OCTOPUS can be modified to facilitate diffusion in organoid culture scaffolds. For example, an orbital shaker can be used to agitate media and generate convective flow in OCTOPUS-containing culture wells as a simple strategy to increase the rate of diffusion, which can contribute to further improving the longevity of organoid models.
Finally, emulating the maturity of native organs in organoid models will require advanced approaches beyond the enhancement of nutrient supply and cell viability demonstrated here to account for the integrated biological complexity of in vivo systems. The assessment is based on the rationale that the short lifespan of organoids is the primary reason for their limited ability to reach later stages of development and acquire mature phenotype. From a developmental biology perspective, however, there is increasing recognition that the limited maturity of organoids in conventional models can also be due to the absence of the surrounding embryonic tissues of developing organs in vivo that provide instructive cues to guide the process of organ development and maturation. Recapitulating this critical aspect of organogenesis in vivo can greatly enhance the ability of OCTOPUS to promote structural and functional maturation of organoids. The results from the co-culture organoid models (
Developing new in vitro technologies for laboratory production and maintenance of organoids is emerging as a major area of investigation in organoid research. Representing this new trend, the work provides a good example of how rational design engineering of conventional organoid culture can advance the ability of organoids to emulate the structural and functional complexity of their in vivo counterparts. By seamlessly integrating engineering novelty into traditional in vitro techniques, the technology offers a simple, practical 3D culture strategy that can be implemented immediately to expand the capabilities of current organoid models. OCTOPUS has the potential for a significant impact on organoid technology and can also provide a powerful platform for various other applications that involve cell and tissue culture in 3D environments.
Investigations into human organoid models will be described in greater detail below.
Having demonstrated the proof-of-concept of OCTOPUS using mouse organoids, the applicability of this technology to human intestinal organoids was explored. To this end, single cell suspension isolated from the small intestine (terminal ileum) of healthy donors were cultured and their self-organization and epithelial differentiation in the disclosed device and Matrigel drops (
Closer examination of the cultured constructs revealed the presence of distinct cell populations and their localized spatial distribution within the developing organoids. For example, proliferative cells identified by positive Ki67 immunostaining were found predominantly at the tip of the buds corresponding to the crypt region (
The enhanced epithelial maturation of enteroids in OCTOPUS was further evidenced by similar trends in the induction of a goblet cell-specific marker (MUC2) (
Recognizing the intrinsic capacity of human intestinal stem cells to give rise to various cell types during organoid development, scRNA-seq analysis was performed to investigate cellular heterogeneity of human enteroids in OCTOPUS. For this study, enteroids were harvested from the disclosed devices at days 7 and 14, and their single-cell transcriptional profiles were examined in comparison to those cultured in Matrigel drops for 7 days—sequencing data from 14-day Matrigel drop culture were excluded in the analysis to avoid confounding factors due to significant cell death observed in this group (
Uniform manifold approximation and projection (UMAP) clustering of the sequencing data obtained from OCTOPUS at day 7 yielded 3 broadly defined groups of cells—absorptive cells, secretory cells, and stem cells—each of which contained multiple subpopulations distinctly identified by the expression of cell-type-specific genes described in previous in vivo studies of the human small intestine (
Importantly, the sequencing results revealed cell populations uniquely present in OCTOPUS. A good example of such cell types is a subset of the absorptive cell population expressing BEST4 (BEST4+ enterocytes) (
To further examine these changes and understand their relevance to the native system, the cellular makeup of the enteroids in OCTOPUS in comparison to the previously published single-cell atlas of the human intestinal epithelium in vivo was analyzed. Results of this analysis showed several differences in epithelial composition between the culture conditions tested in herein. First, extended culture in OCTOPUS led to enrichment of stem cells far beyond what was achievable in Matrigel drop culture as evidenced by their increased proportion in the total population from 7.4% at day 7 to 12.3% by day 14, closely approximating the fraction of intestinal stem cells in vivo (14%) (
Sequencing data also revealed important time- and platform-dependent differences in transcriptional regulation of epithelial maturation. Among the key findings was significantly increased expression of genes specific to mature enterocytes as a result of prolonged culture in OCTOPUS. These genes included i) FABP1, PHGR1, PRAP1, and SLC6A8 for absorptive enterocytes (
Interestingly, in comparison to Matrigel drop culture, OCTOPUS enteroids showed significant downregulation of genes associated with the proliferative capacity of TA cells, such as TOP2A, PCNA, MT1E, and FABP5 (
Finally, single-cell trajectory analysis was performed using Monocle to further characterize the dynamic process of stem cell differentiation during the development of human enteroids in OCTOPUS. When reconstructed in pseudotime on the UMAP plot, the developmental trajectory showed branching into two distinct domains representing absorptive and secretory cell lineages shortly after the initiation of culture (
When combined with the proportion of each cell type, the developmental trajectories allowed for more quantitative characterization and direct comparison of stem cell differentiation in OCTOPUS and Matrigel drops (
Recognizing that in vitro modeling of complex disease is emerging as an active area of investigation in organoid research, efforts shifted to demonstrating the proof-of-principle of using OCTOPUS to construct organoid-based disease models with increased fidelity and physiological relevance. Building upon work on human enteroids, focus moved to modeling human inflammatory bowel disease (IBD), which represents a group of diseases characterized by chronic inflammation of the gastrointestinal tract.
Despite advances in general understanding of IBD, modeling this complex disease remains a significant challenge. Studying IBD often relies on the use of chemical or congenic murine models that require genetic or exogenous manipulations to approximate the phenotype of human IBD. As an alternative approach, researchers have demonstrated conventional 2D and 3D culture of primary or transformed cells (e.g., Caco-2) to generate in vitro analogs of human intestinal tissues that can be subjected to externally applied inflammatory cues. To overcome the limitations of these simplified systems, new efforts are being made to use intestinal organoids to more faithfully model the pathophysiological complexity of IBD. Inspired by this emerging body of work, the feasibility of engineering human enteroids reminiscent of the dysfunctional intestinal epithelium in IBD through long-term culture of patient-derived intestinal organoids in OCTOPUS was explored (
When the cell suspension isolated from the small intestine of IBD patients were seeded into the disclosed device, they underwent self-organization over 7 days into organoids containing budding structures that appeared similar to those observed in normal enteroids derived from healthy donors (
Further analysis using scRNA-seq showed significant alterations reflecting the pathophysiological state of the IBD enteroids. One of the most noticeable changes was a nearly 50% decrease in the proportion of mature enterocytes when compared to normal enteroids (
Having demonstrated the pathophysiological signatures of OCTOPUS-generated IBD enteroids at the molecular and cellular levels, their capacity to recapitulate intestinal abnormalities that develop at the tissue scale during the progression of IBD was then examined. This study focused on intestinal fibrosis, which is a common complication of IBD. The intrinsic ability of the intestine to repair wounds and restore homeostasis can be impaired by repetitive epithelial injury due to chronic inflammation in IBD. Studies have shown that persistent insults can dysregulate tissue-tissue interactions between the intestinal epithelium and the underlying stroma, leading to abnormal remodeling of the subepithelial compartment characterized by hyperproliferation of fibroblasts and excessive deposition of ECM. The goal of the study was to investigate whether the salient features of this pathophysiological fibrogenic process could be recreated in the disclosed IBD organoid model.
To this end, patient-derived IBD enteroids were co-cultured with primary human intestinal fibroblasts in the same hydrogel scaffold to generate a multicellular construct reminiscent of the intestinal epithelium and the underlying stromal tissue in vivo (
These findings match the general patterns of fibrotic tissue remodeling described in previous in vivo studies of the small intestine in IBD patients. Our data also suggest that spontaneous fibrogenesis in the disclosed model is driven by the diseased epithelium of the IBD enteroids. To characterize the soluble pro-fibrotic microenvironment created by these epithelial cells, the production of transforming growth factor (TGF)-β1, which is the member of the TGF-β superfamily overexpressed by the intestinal epithelium in IBD that selectively activates ECM synthesis by mesenchymal cells, was measured. As expected, TGF-β1 production was significantly upregulated in the IBD model compared to normal enteroids (
With rapid progress in organoid technology, there is a significant increasing demand for advanced organoid models capable of emulating more complex structure and physiological function of native organs. As a representative example, integrating vasculature into organoid cultures is emerging as an area of increasing interest in ongoing research efforts to advance the capabilities and potential of organoid technology. Vascularization of organoids is necessary for mimicking vascularity of native tissues and vascular contributions to parenchymal function but it has also been suggested as a promising strategy to improve nutrient and oxygen supply in 3D culture for enhanced organoid growth and maturation. The process of generating vascularized organoids and perfusing them in a controlled manner, however, is prohibitively complex and often requires specialized techniques and culture systems not easily accessible to non-engineers.
Motivated by this problem, advanced prototype of OCTOPUS that provides new capabilities to engineer vascularized, perfusable human organoids while still offering the simplicity and convenience of the original platform was created (
For proof-of-concept demonstration, we seeded OCTOPUS-EVO with a mixture of fibrin and Matrigel precursors containing human intestinal cells, endothelial cells, and fibroblasts. This co-culture system supported rapid self-organization of stem cells into enteroids within 2-3 days of culture (
Building upon the demonstration of IBD enteroids (
Finally, the observation of endothelial activation in the vascularized IBD enteroids led to investigation of whether vascular perfusability of the dislcosed model could be exploited to simulate the recruitment of blood-borne immune cells in IBD. In vivo evidence has established a marked increase in the recruitment of circulating blood monocytes to the intestinal mucosa as one of the key immunological events during the development of IBD. Indeed, the disclosed IBD model perfused with human peripheral blood monocytes showed a large number of cells in the enteroid-associated blood vessels, as well as in the lumen of the enteroids (
This data altogether provide the proof-of-concept of OCTOPUS-EVO and demonstrate its potential as an accessible in vitro platform to engineer vascularized, perfusable organoids that can expand the capabilities of conventional organoid cultures.
In response to the increasing need for new technologies for organoid research, the present disclosure describes a microengineered platform to reconfigure the three-dimensionality of conventional organoid culture. OCTOPUS provides a simple yet effective means to address the problem of limited nutrient supply inherent in 3D culture. By enabling controlled production of open 3D culture scaffolds with significantly decreased thickness, this system serves to reduce the distance and spatial variability of nutrient and oxygen diffusion to growing organoids. In comparison to conventional Matrigel drop culture, this design makes it possible to engineer a more uniform, unrestricted soluble microenvironment beneficial for long-term culture of organoids. The improved mass transport characteristics due to significantly reduced diffusion limitations also decrease the effective culture volume of the disclosed system, which is an inverse measure of the ability of cells to process and control their environment during culture. As a result, stem cells and organoids in OCTOPUS have better control over their local microenvironment during development. Data described herein show that these desirable features of OCTOPUS can increase the size and maturity of organoids beyond what is achievable using conventional techniques and may enable the production of more realistic multicellular constructs for in vitro modeling of organogenesis and disease development.
OCTOPUS enables uninterrupted, continuous organoid culture for extended periods of time. As shown by scRNA-seq of the human enteroid model, doubling the duration of uninterrupted culture using OCTOPUS greatly promoted enterocyte differentiation in organoids to generate a more physiological intestinal epithelium that contained substantially larger numbers of functionally mature enterocytes, as compared to Matrigel drop.
Although the ability to support continuous long-term culture is a key advantage of OCTOPUS, the data herein reveal additionally desirable features of organoid development in the disclosed system. After 7 days of culture, for example, the size of intestinal organoids and the expression of virtually every marker of epithelial maturation were significantly greater in OCTOPUS. ScRNA-seq analysis provided further evidence that human enteroids in OCTOPUS more faithfully recapitulated the cellular heterogeneity of the native intestinal epithelium, as well as the relative abundance of differentiated cell types and their physiological gene expression profiles, when compared to those cultured in conventional Matrigel drops for the same amount of time. These results suggest that OCTOPUS is capable of accelerating the growth and maturation of organoids at the early stage of development.
By leveraging these capabilities, this work demonstrated the feasibility of developing a specialized organoid model that can recapitulate morphological, functional, and transcriptional characteristics of the diseased human intestinal epithelium in IBD. Interestingly, many of the pathophysiological alterations observed in this model did not occur in Matrigel drop culture of patient-derived organoids. Another observation of the OCTOPUS IBD model was increased expression of several long noncoding RNAs (lncRNAs). This finding may have important implications for the emerging investigation of lncRNA biology in IBD. Recent evidence suggests active involvement of lncRNAs in mediating key disease processes of IBD associated with epithelial permeability, apoptosis, and inflammation. To this end, the scRNA-seq data reveal a set of lncRNAs that have not been implicated in IBD. LINC02159 and LINC02577 are among these genes that have been shown to play a role in tumorigenesis by promoting the proliferation of colorectal cancer cells. LINC01210 is another lncRNA previously described as a regulator of colorectal and ovarian cancer cell proliferation and invasion.
Inclusion of intestinal fibroblasts in this model permitted in vitro reproduction of intestinal fibrosis. Unlike previous demonstrations of organoid-based fibrosis models generated by treatment with exogenous fibrogenic factors (e.g., TGF-β), the disclosed co-culture system spontaneously developed fibrosis without external input to recapitulate the key features of abnormal matrix remodeling described in the small intestine of IBD patients. This finding supports the general notion of the diseased or persistently injured epithelium as the driver of pathophysiological organ fibrosis that can activate effector cells in the subepithelial compartment. Thus, the disclosed system may provide a simple yet enabling platform for organoid-based mechanistic investigation of dysregulated fibrogenesis in the intestine. Given that the biological processes underlying the development of fibrosis are conserved across organs, the same device and organoid culture techniques may be applicable to studying fibrotic diseases in other organs.
The demonstration of organoid vascularization highlights the advanced capabilities and potential of OCTOPUS. OCTOPUS-EVO enabled the concurrent, spontaneous process of organogenesis and vasculogenesis in the same culture scaffold to produce vascularized, perfusable human enteroids that can recreate the vascular-parenchymal interface and more complex physiological responses of native organs. Researchers have recently introduced techniques for organoid vascularization, including in vivo transplantation of organoids into vascular-rich organs such as the brain, kidney, lung, and pancreas, but generating such constructs with controlled vascular perfusion in vitro remains a major challenge. OCTOPUS-EVO provides an accessible means to tackle this challenge and increase the complexity of organoid models at the convenience and simplicity of conventional 3D culture without requiring specialized engineering systems. Vascularized enteroids in the disclosed device had significantly larger size compared to non-vascularized ones, supporting the notion that organoid vascularization is a promising strategy to facilitate organoid growth. Presumably, vascularization of the culture scaffold increases nutrient and oxygen supply to permit more efficient and rapid organoid development. Based on a large body of evidence demonstrating endothelial interactions with parenchymal tissues, it is also possible that biological crosstalk between the vasculature and organoids may be responsible for increased organoid growth.
While OCTOPUS represents considerable changes to the design of traditional organoid models, the implementation of this system does not require any modification of established culture protocols and workflow, nor does it rely on specialized equipment or personnel. Essential to this advantage is the design of OCTOPUS as a ready-to-use and easily-accessible culture insert that is directly compatible with standard well plates and laboratory infrastructure. As exemplified by the disclosed intestinal models, generating mature organoids in OCTOPUS can readily be accomplished in traditional laboratory settings based on materials and experimental procedures commonly used in conventional techniques. This is an important aspect of the disclosed methods that makes OCTOPUS an immediately deployable and readily accessible culture platform, which may contribute to rapid dissemination of the technology for widespread use.
The below described methods were applied, as appropriate, for each of the above-described Examples.
For organoid cultures, cryopreserved mouse intestinal organoids (70931, STEMCELL Technologies, Canada) and cryopreserved mouse hepatic progenitor organoids (70932, STEM-CELL Technologies, Canada) were used. Intestinal and liver organoids were cultured in 24-well plates according to the manufacturer's protocols using IntestiCult™ organoid growth medium (06005, STEMCELL Technologies, Canada) and Hepaticult™ organoid growth medium (06030, STEM-CELL Technologies, Canada), respectively. To briefly explain, previously existing Matrigel drop was dissolved by incubating in Dispase. Upon incubation for 30 minutes, organoids were physically dissociated into single cell suspension and then transferred to a 15 ml falcon tube and centrifuged at 290×g to obtain stem cell pellet. 100 μl of complete organoid growth medium was then added to the pellet. After 100 μl of cold Matrigel was added, the suspension was gently pipetted up and down 10 times for thorough mixing. Using a pre-wetted 200 μl tip, 50 μl of the organoid/Matrigel mixture was injected into a 24-well plate to form Matrigel drop. The drop-containing well plates were then incubated at 37° C. and 5% CO2 for 10 minutes to allow gelation of Matrigel. Upon completion of this step, 750 μl of pre-warmed organoid growth medium was added to each well. Organoids were passaged every 5-7 days in fresh Matrigel until use as recommended by the manufacturer.
Regarding human enteroid lines, enteroid lines generated from terminal ileum were provided by the Children's Hospital of Philadelphia Gastrointestinal Epithelium Modeling Program under an Institutional Review Board-approved protocol (13042). All parents of patients provided written informed consent. Enteroid lines were generated. Briefly, two biopsy tissue fragments were rinsed 3 times in 1 ml cold sterile PBS, then incubated in cold chelation buffer for 30 minutes on a turntable in a cold room, followed by mechanical dissociation (scraping) of epithelial layer. The fragments were strained through a 100 μm strainer to deplete the villi and resuspended in 80% Matrigel, then seeded at the density of crypts per 30 μL drop. The droplets solidified at 37° C. for 30 minutes, and 500 μl human IntestiCult (STEMCELL Technologies; complete when supplemented with Penicillin-Streptomycin (Gibco)) was added per well. Y-27632 (SelleckChem; final conc. 10 μM) was added to culture medium at seeding only.
Regarding maintenance and passage of human enteroids, enteroid media was changed three times per week. On day 14, the cultures were passaged and/or cryopreserved in CryoStor CS-10 (STEMCELL Technologies). To passage, the Matrigel droplet was dislodged by pipetting up and down through a P1000 tip and transferred into a 1.5 ml microfuge tube, followed by centrifugation and washing with ice-cold HBSS. Enteroids were mechanically dissociated into fragments by pipetting 10 times through a P200 tip placed on top of a P1000 tip, followed by centrifugation. The pellet was reconstituted in 80% Matrigel and seeded as 30 μl drops at a split ratio of 1:4. Subsequent cultures are ready for passage and/or cryopreservation on day 7.
Regarding formation of 3D organoid constructs in OCTOPUS, standard 24-well plates containing OCTOPUS inserts were first sterilized by exposure to ultraviolet (UV) light (Electro-lite ELC-500) for 30 minutes. Subsequently, the culture chambers in OCTOPUS were filled with 2 mg/ml (w/v in 10 mM Tris-HCl buffer, pH 8.5) of dopamine hydrochloride solution at room temperature (RT) for 2 hours to form a surface coating for enhanced adhesion of Matrigel to PDMS. The poly(dopamine) (PDA)-treated devices were kept sterile until use. To form organoids in the disclosed device, the pellets were made first. To this end, existing Matrigel was dissolved by incubating Matrigel drop in Dispase. Cells were then transferred to a 15 mL falcon tube and centrifuged at 290×g to obtain a stem cell pellet. Then, 100 μl of complete IntestiCult™ organoid growth medium was added to the pellet. After 100 μl of cold Matrigel was added, the suspension was gently pipetted up and down 10 times for thorough mixing. For human intestinal enteroids, a cell pellet can be resuspendeded in 80% Matrigel. Using a pre-wetted 200 μl tip, 100 μl of the organoid/Matrigel mixture was injected into OCTOPUS through the injection port. The OCTOPUS-containing well plates were then incubated at 37° C. and 5% CO2 for 10 minutes to allow gelation of Matrigel. Upon completion, 750 μl of pre-warmed IntestiCult™ organoid growth medium was added to each well. The OCTOPUS plates were maintained in cell culture incubators at 37° C. and 5% CO2. During long-term culture, media exchange was conducted every other day.
For measurement and quantification of caspase-3, annexin V, TNFa, TGFβ-1, IL-6 and IL-8 in the human enteroids, conditioned media were collected on day 14 of culture and analyzed using cleaved caspase-3 (Asp175) ELISA kit (ab220655, abcam), human annexin V ELISA kit (ab223863, abcam), human TNF alpha ELISA kit (ab181421, abcam), human TGF beta 1 ELISA kit (ab100647, abcam), human IL-6 ELISA kit (ab178013, abcam), and human IL-8 ELISA kit (ab214030, abcam). Each assay was performed following the manufacturer's protocol. Briefly, 100 μl of a standard solution or sample media was added to each well. After 2-hour incubation, the well was washed 5 times with 300 μl of manufacturer-provided wash buffer and incubated with secondary antibody for 1 hour. After washing, 100 μl of TMB substrate was added to each well and incubated for 20 minutes in the dark. Finally, 100 μl of stop solution was added to each well, and the plate was measured in a plate reader (M200, Tecan, Switzerland). For all ELISA assays, we used a multimode plate reader (M200, Tecan, Switzerland) to measure the optical density of samples. A standard curve was generated by plotting the mean optical density and concentration for each standard using a four-parameter logistic curve fitting method. Sample measurements were converted to target concentrations using the standard curves.
To model intestinal fibrosis as a complication of IBD, human intestinal stem cells were co-cultured with 1×106 cells/ml of primary human intestinal fibroblasts in Matrigel (356255, Corning, USA). This cell-containing hydrogel solution was injected into the device to form microtissue constructs in the organoid culture chambers. After gelation for 15 minutes in a regular cell culture incubator, 750 μl of IntestiCult™ organoid growth medium (06010, STEMCELL Technologies, Canada) was added into each well and maintained for 14 days to induce intestinal organoid development and fibroblast proliferation. During this period, the media were replenished every other day.
Regarding the formation of vascularized human enteroids in OCTOPUS-EVO, the fully assembled device was sterilized before cell culture by exposing it to ultraviolet (UV) light (Electro-lite ELC-500) for at least 30 minutes. To engineer vascularized organoids in OCTOPUS-EVO, 20 μl of cell suspension solution containing fibrinogen (5 mg/ml; F8630, Sigma), thrombin (1 U/ml; T7513, Sigma), aprotinin (0.15 U/ml; A1153, Sigma), human intestinal stem cells, primary human umbilical vein endothelial cells (HUVECs) (5×106 cells/ml), and primary normal human lung fibroblasts (NHLFs) (1×106 cells/ml) was prepared and injected it into the open cell culture chamber through its inlet access port. The device was then left in a cell culture incubator at 37° C. and 5% CO2 for 30 minutes. Upon gelation, IntestiCult media mixed with EGM-2 endothelial media were added to the medium reservoirs and the side microchannels. Following the formation of cell-laden hydrogel construct, the side microchannels were incubated with a fibronectin solution (25 μg/ml in PBS; 356008, Corning) for 2 hours at 37° C. to create ECM coating on the channel surface. Then, the channels were washed once with IntestiCult/EGM-2, and 10 μl of HUVEC suspension (1×107 cells/ml) was introduced into both channels. The seeded cells were allowed to attach to the channel surface over a period of 1 hour. Upon 1 hour incubation, pre-warmed media was added to each medium reservoir. This culture condition allowed the endothelial cells to form confluent monolayers on the surface of the side channels and the hydrogel scaffold to induce anastomosis between the endothelial lining and the self-assembled vasculature in the hydrogel.
To examine the viability of organoids, Live/Dead™ Viability/Cytotoxicity Kit was used for mammalian cells (L3224, ThermoFisher Scientific, USA). For this assay, a mixture of calcein AM (2 μM) and ethidium homodimer-1 (4 μM) in live-cell imaging solution was introduced into the OCTOPUS-containing wells and incubated at RT for 30 minutes. Subsequently, the samples were washed with phosphate-buffered saline (PBS) three times, after which the labeled cells were examined using a laser scanning confocal microscope (LSM 800, Carl Zeiss, Germany). For quantitative analysis, the fraction of healthy and necrotic organoids was calculated from fluorescence generated by calcein AM and ethidium homodimer-1, respectively. In each device, 30 organoids were used for the analysis.
To examine the spatiotemporal patterns of diffusion in OCTOPUS and hydrogel drops, FITC-dextran, either 4 kDa FITC-dextran or 70 kDa FITC-dextran (FD70S-100MG, Sigma, USA), was used as a fluorescent tracer for visualization. For this assay, the organoid culture medium was replaced with a FITC-dextran solution (50 μg/ml in PBS). Dextran diffusion was monitored and visualized using a laser scanning confocal microscope (LSM 800, Carl Zeiss, Germany). Time-lapse images were acquired for 120 minutes and processed using ZEN software (Zeiss, Germany) to measure temporal changes in fluorescence intensity at defined locations within the hydrogel scaffolds.
To detect proliferating cells within the intestinal organoids, EdU assay/EdU staining proliferation kit-iFluor 647 (ab222421, abcam, USA) was used. Briefly, the organoids were incubated with a EdU solution (20 μM in medium) for 3 hours under normal culture conditions (5% CO2 at 37° C.). The organoids were then washed twice with PBS, fixed in 4% formaldehyde, and permeabilized using a permeabilization buffer, according to the manufacturer's protocol. The samples were stained with iFluor 647 azide dye and visualized using a confocal microscope (LSM 800, Carl Zeiss, Germany).
For calcium imaging, the organoid media were removed from the culture wells, and the organoid constructs were washed once in live-cell imaging solution (LCIS). The organoids were then loaded with Fluo-4 calcium imaging solution (F10489, ThermoFisher Scientific, USA) prepared according to the manufacturer's protocol. The samples were incubated at 37° C. for 30 minutes, which was followed by another 30-minute incubation at room temperature. Subsequently, the Fluo-4 solution was removed, and the organoids were washed once with LCIS. All samples were kept in fresh LCIS until use. An inverted epi-fluorescence microscope (Axio Observer D1, Zeiss, Germany) was used to visualize calcium staining of organoids upon stimulation with 100 μM of ATP (A1852, Sigma, USA) and 50 mM of glucose (G7021, Sigma, USA).
For ratiometric analysis of changes in Ca2+ levels, fluorescence intensity for each or-ganoid was measured during an experiment, and values were normalized by their resting intensities using the equation below.
ΔCa2+=(F−Frest)/Frest (1)
To analyze GLP-1 and mucin 2 secretion from the intestinal organoids, the media in the wells were collected on days 5, 7, and 10 of culture. Multi-species GLP-1 total ELISA kit (EZGLP1T-36K, Millipore Sigma, USA), Glucagon-like peptide-1 (active) ELISA kit (EGLP-35K, Millipore Sigma, USA), and MUC2 ELISA kit (ABIN6730976, antibodies-online Inc, USA) were used to measure the concentrations of GLP-1 total, GLP-1 active, and mucin 2, respectively. Each assay was performed following the manufacturer's protocol. Briefly, 100 μl of a standard solution or sample media was added to each well. After 2-hour incubation, the well was washed 5 times with 300 μl of manufacturer-provided wash buffer and incubated with secondary antibody for 1 hour. After washing, 100 μl of TMB substrate was added to each well and incubated for 20 minutes in the dark. 100 μl of stop solution was added to each well, and the plate was measured in a plate reader (M200, Tecan, Switzerland).
For the analysis of fibronectin production in the intestinal fibrosis model, a mouse fibronectin ELISA kit (ab108849, abcam, USA) was used. The media in the wells were collected at specified time points and assayed using manufacturer-provided protocols. First, 50 μl of standard or device-collected samples were added into each well and incubated for 2 hours at room temperature. Subsequently, the wells were washed 5 times with 300 μl of wash buffer solution and then incubated with fibronectin antibody for 1 hour. After washing, the streptavidin-peroxidase conjugate was added to each well, incubated for 30 minutes, and washed again. The samples were incubated with 50 μl of chromogen substrate for 10 minutes, followed by the introduction of 50 μl of stop solution.
For all ELISA assays, a multimode plate reader (M200, Tecan, Switzerland) was used to measure the optical density of samples. A standard curve was generated by plotting the mean optical density and concentration for each standard using a four-parameter logistic curve fitting method. Sample measurements were converted to target concentrations using the standard curves.
For certain co-culture demonstrations, primary mouse intestinal fibroblasts (mIFs) and primary human umbilical vein endothelial cells (HUVECs) were used. For initial expansion from cryostorage, mIFs and HUVECs were cultured in 75 cm2 flasks according to the manufacturer's protocols using complete fibroblast medium (M2267, Cell Biologics, USA) and endothelial cell growth medium (EGM)-2 (CC-3162, Lonza, Switzerland) supplemented with growth factors, respectively. Primary mIFs and primary human intestinal fibroblasts were used for modeling intestinal fibrosis. All cells were between passages 3 and 6.
To form an intestinal fibrosis model suitable for, e.g. drug testing, in OCTOPUS, mouse intestinal stem cells were mixed with 1×106 cells/ml of mouse intestinal fibroblasts in Matrigel (356255, Corning, USA). This cell-containing hydrogel solution was injected into the device to form microtissue constructs in the organoid culture chambers. After gelation for 15 minutes in a regular cell culture incubator, 750 μl of IntestiCult™ organoid growth medium (06005, STEMCELL Technologies, Canada) was added into each well and maintained for 5 days to induce intestinal organoid development and fibroblast proliferation. During this period, the media were replenished every other day. To induce fibrosis, 1 ng/ml of TGF-β (T5050, Sigma, USA) was added to the culture wells on day 5 and maintained for additional 7 days. Drug administration occurred on day 12 using commercially available Pirfenidone (P1871, TCI America, USA) and Nintedanib (S1010, Selleckchem, USA) at specified concentrations. The fibrosis model was treated with drugs for 48 hours, after which changes in its fibrotic phenotype were analyzed using the methods described herein.
To examine oxygen diffusion in OCTOPUS and Matrigel drops, Dichloritis(1,10-phenanthroline)ruthenium(II) hydrate (Ru(phen)3) (Sigma, Cat. #343714) was used as an oxygen indicator—dissolved oxygen molecules induce quenching of this fluorescent dye. Briefly, 15 μl of Ru(phen)3 (2 mM) and 270 μl of Matrigel were mixed with 15 μl of sodium sulfite (200 mM) (Sigma, Cat. #S0505), which was used to remove remaining aqueous oxygen in Matrigel. The mixture was then used to generate 3D tissue constructs in OCTOPUS and Matrigel drop. Oxygen diffusion was monitored by confocal microscopy, during which images were acquired at defined time intervals and locations within the constructs. The captured images were analyzed using ImageJ to measure spatiotemporal changes in fluorescence intensity.
To test the perfusability of the microengineered vascular network, fluorescently labeled 1-μm microbeads (FluoSpheres; F-8815, ThermoFisher) were used as flow tracers. To generate flow through the vasculature, media in the reservoirs was aspirated and a bead solution was inserted into one of the side microchannels. This configuration created a gradient of hydrostatic pressures across the hydrogel scaffold and provided driving force for the flow of microbeads through the vessels. Vascular perfusion was monitored and visualized using a laser scanning confocal microscope (LSM 800, Carl Zeiss, Germany).
To perform a monocyte infiltration assay, human peripheral blood monocytes were obtained from the Human Immunology Core at the University of Pennsylvania. To test endothelial adhesion of monocytes in the disclosed system, cells were labeled with a fluorescent dye (CellTracker Deep Red, ThermoFisher) and suspended in IntestiCult/EGM-2 media at the final concentration of 3×106 cells/ml. The cells were then injected into the vessels through one of the side microchannels and allowed to flow through the vasculature for 24 hours in a cell culture incubator. At the completion of perfusion, the device was washed with DPBS three times and examined to analyze the number of adhered, transmigrated, and infiltrated monocytes.
Atomic force microscopy (AFM, MFP-3D-BIO, Asylum) was used to measure the stiffness of hydrated microtissues in the intestinal fibrosis models. A gold-coated cantilever (SCONT tip, NANOSENSORS) with a spring constant of 14.58 pN/nm and a pyramid indenter was used to obtain force-indentation curves. The tissue samples in the open chambers were used directly without any modification. For the AFM measurement, the OCTOPUS insert containing microtissue was removed from the plate and mounted on the instrument. After wetting the microtissue with a drop of PBS, its mechanical property was measured with the scanning probe. Young's modulus was calculated from the force indentation data using the Atomic J software.
For immunofluorescence staining, cells in OCTOPUS were washed with PBS twice, fixed with 4% paraformaldehyde (Electron Microscopy Sciences, USA) for 15 minutes at room temperature, and washed again twice with PBS. The cells were then permeabilized with 0.1% Triton X-100 (Sigma) in PBS for 3 minutes and exposed to blocking buffer composed of PBS and 3% bovine serum albumin (BSA; Sigma) for overnight at 4° C. After washing with PBS twice, the cells were immunostained for actin filaments (Phalloidin-iFluor 488 Reagent, ab176753, 1:1000, abcam, USA; Phalloidin-iFluor 594 Reagent, ab176757, 1:1000, abcam, USA), mature epithelial cells (anti-EPCAM antibody, ab71916, 1:250, abcam, USA; anti-HNF-4-alpha antibody [K9218]-ChIP Grade, ab41898, 1:500, abcam, USA), stem cells (anti-Ki67 antibody, ab15580, 1:1000, abcam, USA; Lgr5 monoclonal antibody, MA5-25644, 1:1000, ThermoFisher Scientific, USA), enterocytes (anti-Villin antibody [3E5G11]-N-terminal, ab201989, 1:500, abcam, USA), goblet cells (anti-MUC2 antibody, ab90007, 1:200, abcam, USA), enteroendocrine cells (anti-Somatostatin antibody [M09204], ab30788, 1:100, abcam, USA), peptide transporter 1 (anti-SLC15A1/PEPT1 antibody, ab203043, 1:100, abcam, USA), glucose transporter 1 (anti-Glucose transporter GLUT1 antibody [SPM498], ab40084, 1:250, abcam, USA), endothelial cells (anti-CD31 antibody [JC/70A] (Alexa Fluor® 488), ab215911, 1:100, abcam, USA), alpha smooth muscle actin (recombinant anti-alpha smooth muscle Actin antibody [E184], ab32575, 1:500, abcam, USA), fibronectin (anti-Fibronectin antibody [IST-9], ab6328, 1:200, abcam, USA), alpha smooth muscle actin (recombinant anti-alpha smooth muscle Actin antibody [E184], ab32575, 1:500, abcam, USA), fibronectin (anti-Fibronectin antibody [IST-9], ab6328, 1:200, abcam, USA), cleaved caspase-3 (anti-cleaved caspase-3 antibody [E83-77], ab32042, 1:200, abcam, USA), annexin V (anti-annexin V/ANXA5 antibody [EPR3980], ab108194, 1:500, abcam, USA), or ICAM1 (anti-ICAM1 antibody [EPR24639-3], ab282575, 1:500, abcam, USA). After overnight incubation with primary antibody at 4° C., the cells were washed twice with PBS and incubated with secondary antibody (Goat anti-Rabbit IgG H&L (Alexa Fluor® 488), ab150077, 1:1000, abcam, USA; Goat anti-Mouse IgG H&L (Alexa Fluor® 488), ab150113, 1:1000, abcam, USA; Goat anti-Mouse IgG H&L (Alexa Fluor® 594), ab150116, 1:1000, abcam, USA; Goat anti-Rabbit IgG H&L (Alexa Fluor® 594), ab150080, 1:1000, abcam, USA) overnight at 4° C. For nuclear staining, DAPI (D1306, Ther-moFisher Scientific, USA) diluted at 1:1000 was used. Fluorescence images of the stained cells were acquired using a laser scanning confocal microscope (LSM 800, Carl Zeiss, Germany) and processed using ZEN software (Zeiss, Germany) and ImageJ software.
For hematoxylin and eosin (H&E) staining of human enteroid, organoids were washed with cold PBS and fixed with 4% paraformaldehyde (Electron Microscopy Sciences, USA). The organoids were then resuspended in embedding gel composed of 2% bacto-agar and 2.5% gelatin and transferred as a droplet onto the embedding rack. After the gel was solidified for 30 minutes, the organoid embedded gel was placed in the pre-labeled tissue cassette and submerged in 70% ethanol. The slides containing paraffin sections were deparaffinized and rehydrated by immersing the slides sequentially into 3× Xylene, 2×100% ethanol, 95-95-80-70% ethanol, and distilled water. Then, the slides were immersed in 10 mM citric acid buffer (pH 6.0) and incubated in microwave oven for 15 minutes. After gently rinsing the slides, tissue sections were blocked with protein blocking agent. To perform H&E staining, the slides were immersed in Hematoxylin followed by rinsing with deionized water. The slides were further immersed in Eosin for 30 seconds and dehydrated in 95% ethanol-100% ethanol-xylene solutions. Tissue sections were covered with coverslip slides using Permount and stored until analysis.
Quantitative RT-PCR analysis was performed as follows. For RNA isolation, organoids were harvested by dissolving Matrigel including organoids with cold PBS. Following centrifugation at 300×g for 5 minutes at 4° C., the supernatant was removed and the pelleted organoids were resuspended in 350 μL of RLT buffer (QIAGEN). Total RNA was isolated using the RNeasy Mini Kit (QIAGEN) according to the manufacturer's instructions. cDNA was synthesized using iScript cDNA Synthesis Kit (Bio-Rad) following the manufacturer's instructions. Quantitative RT-PCR was performed using TaqMan® gene expression assays.
For single-cell sequencing analysis, collected organoids were incubated in trypsin for 10 minutes at 37° C. and passed through a 20-μm cell strainer. Isolated single cells were re-suspended at a density of 700 live cells/μl in DMEM with 5% fetal bovine serum (FBS). The cells were then stained with trypan blue to check their viability and counted under the microscope twice to determine the average cell concentration.
Single-cell suspension for each organoid sample was loaded onto a separate channel of a Chromium 10× Genomics Single Cell 3′ Reagent Kit v2 library chip (10× Genomics) according to the manufacturer's protocol. RNA transcripts from single cells were uniquely barcoded and reverse-transcribed. cDNA sequencing libraries were prepared according to the manufacturer's protocol (10× user guide for library prep) and sequenced on an Illumina NovaSeq 6000 using an S1 100 cycles flow cell v1.5. Library quality control was done using Agilent TapeStation for sizing (bp) and KAPA qPCR for concentration (nM). Raw sequence reads data were processed using the CellRanger pipeline (10× Genomics, v.5.0.0) for demultiplexing and aligned to the human genome GRCh38 transcriptome. Sample data was aggregated using the CellRanger aggr pipeline and libraries were normalized for sequencing depth across the sample set. A total of 5 organoid sample count matrices were merged together for cell type identification and direct comparisons.
Regarding statistical analysis, the sample size for each experiment was determined on the basis of a minimum of n=3 independent devices for each experimental group. Data were analyzed with Student's t-test using OriginLab (OriginLab Corporation, USA) and presented as mean±S.E.M. Statistical significance of the obtained data was attributed to values of *P<0.05, **P<0.01, and ***P<0.001 as determined by one-way ANOVA analysis.
The following Embodiments are illustrative only and do not limit the scope of the present disclosure or the appended claims.
Embodiment 1. A device for culturing organoids, comprising: an access port configured to receive a solution; a loading chamber, wherein the access port is located in the loading chamber; and a plurality of culture chambers, wherein the culture chambers are radiated from the loading chamber so that the solution injected into the loading chamber through the access port is distributed into the plurality of culture chambers, wherein the plurality of culture chambers are open to an external environment and comprises a protruding edge at an opening of the plurality of culture chambers.
Embodiment 2. The device of Embodiment 1, wherein the device comprises poly(dimethyl siloxane).
Embodiment 3. The device of Embodiment 1 or 2, wherein the device is optically transparent.
Embodiment 4. The device of any one of Embodiments 1-3, wherein the access port is located in a center of the loading chamber.
Embodiment 5. The device of Embodiment 4, wherein the plurality of culture chambers are symmetrical with respect to rotations about the access port.
Embodiment 6. The device of Embodiment 5, wherein the solution injected into the loading chamber through the access port is evenly distributed into the plurality of culture chambers.
Embodiment 7. The device of any one of Embodiments 1-6, wherein the device is configured to contact a culture media from the external environment through the opening of the plurality of culture chambers.
Embodiment 8. The device of Embodiment 1, wherein the solution is a hydrogel solution.
Embodiment 9. The device of Embodiment 1, wherein the hydrogel solution comprises cells or organoids.
Embodiment 10. The device of Embodiment 1, wherein the organoids are human organoids.
Embodiment 11. The device of any one of Embodiments 1-10, wherein each of the culture chambers has a width or a height ranging from about 100 μm to about 5 cm.
Embodiment 12. The device of Embodiment 11, wherein each of the culture chambers has a width and a height of about 1 cm.
Embodiment 13. The device of any one of Embodiments 1-12, wherein at least about 80% of the organoids in the culture chamber are viable at day 21 of culturing.
Embodiment 14. The device of any one of Embodiments 1-13, wherein the protruding edge is configured to pin a meniscus of the solution at the opening of the culture chambers, allowing filling of the culture chambers without spillage of the solution through the opening.
Embodiment 15. The device of any one of Embodiments 1-14, wherein each culture chamber comprises a different type of cells or organoids for co-culturing.
Embodiment 16. The device of any one of Embodiments 1-15, wherein growth of the organoids continues for at least about 21 days.
Embodiment 17. The device of any one of Embodiments 1-16, wherein a size of the organoids increases for at least about 21 days.
Embodiment 18. The device of Embodiment 17, wherein the device decreases variability in the size of the organoids.
Embodiment 19. A method for culturing organoids, comprising: injecting a solution including cells or organoids into a loading chamber through an access port; filling a plurality of culture chambers with the solution including cells or organoids, wherein the culture chambers are radiated from the loading chamber so that the solution injected into the loading chamber is distributed into the plurality of culture chambers, wherein the plurality of culture chambers are open to an external environment and comprises a protruding edge at an opening of the culture chambers for preventing spillage of the solution through the opening; and providing a culture media to the device through the opening of the plurality of culture chambers.
Embodiment 20. The method of Embodiment 19, wherein the access port is located in a center of the loading chamber.
Embodiment 21. The method of Embodiment 20, wherein the plurality of culture chambers are symmetrical with respect to rotations about the access port.
Embodiment 22. The method of Embodiment 21, wherein the solution injected into the loading chamber through the access port is evenly distributed into the plurality of culture chambers.
Embodiment 23. The method of any one of Embodiments 19-22, wherein the solution is a hydrogel solution.
Embodiment 24. The method of any one of Embodiments 19-23, wherein the organoids are human organoids.
Embodiment 25. The method of Embodiment 23 or 24, wherein the hydrogel solution is solidified to form a hydrogel in the plurality of culture chambers after being injected into the loading chamber and distributed into the plurality of culture chambers.
Embodiment 26. The method of any one of Embodiments 19-25, wherein at least about 80% of the organoids in the culture chamber are viable at day 21 of culturing.
Embodiment 27. The method of any one of Embodiments 19-26, wherein each culture chamber comprises a different type of cells or organoids for co-culturing.
Embodiment 28. The method of any one of Embodiments 19-27, wherein growth of the organoids continues for at least about 21 days.
Embodiment 29. The method of any one of Embodiments 19-28, wherein a size of the organoids increases for at least about 21 days.
Embodiment 30. The method of Embodiment 17, wherein the device decreases variability in the size of the organoids.
Embodiment 31. The method of any one of Embodiments 19-30, wherein the culture media comprises soluble factors.
Embodiment 32. The method of Embodiment 31, wherein the soluble factors are selected from the group consisting of a growth factor, an active agent, and a combination thereof.
Embodiment 33. The method of any one of Embodiments 19-32, further comprising maturing the organoids.
Embodiment 34. The method of any one of Embodiments 19-33, further comprising assessing viability and maturation of the organoids in the plurality of culture chambers.
All patents, patent applications, publications, product descriptions, and protocols, cited in this specification are hereby incorporated by reference in their entireties. In case of a conflict in terminology, the present disclosure controls.
While it will become apparent that the subject matter herein described is well calculated to achieve the benefits and advantages set forth above, the presently disclosed subject matter is not to be limited in scope by the specific embodiments described herein. It will be appreciated that the disclosed subject matter is susceptible to modification, variation, and change without departing from the spirit thereof. Those skilled in the art will recognize or be able to ascertain using no more than routine experimentation, many equivalents to the specific embodiments described herein. Such equivalents are intended to be encompassed by the following claims.
This application claims the benefit of U.S. Patent Application No. 63/121,684, filed on Dec. 4, 2020, the content of which is hereby incorporated by reference in its entirety.
This invention was made with government support under HL127720 awarded by the National Institutes of Health and 1548571 awarded by the National Science Foundation. The government has certain rights in the invention.
Filing Document | Filing Date | Country | Kind |
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PCT/US2021/072762 | 12/6/2021 | WO |
Number | Date | Country | |
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63121684 | Dec 2020 | US |