Scalable and physiologically relevant microenvironments for human pluripotent stem cell expansion and differentiation

Human pluripotent stem cells (hPSCs), including human embryonic stem cells (hESCs) and induced pluripotent stem cells (iPSCs), are attractive cell sources for various biomedical applications including cell therapies [1], tissue biofabrication [2–4], drug screening and toxicity tests [5, 6]. These applications require large numbers of high quality cells [7]. For instance, it is estimated that ∼10⁹ cardiomyocytes, 10⁹ β cells, 10¹² red blood cells, and 10¹¹ platelets are needed for treating one myocardial infarction or diabetic patient, or one blood or platelet transfusion, respectively [1]. Approximately 10¹⁰ cells are required for engineering a human-size heart or liver [5], or for screening a library with one million compounds once [6]. Considering the large patient population and the large numbers of chemical libraries that can be screened, massive numbers of cells are therefore needed [7].

However, the scalable and cost-effective culturing of high quality hPSCs and their derivatives, especially for clinical applications, remains a challenge [1, 7, 8]. In vivo, majority of human cells including the hESCs reside in 3D microenvironments that have plenty of cell–cell and cell–ECM (extracellular matrix) interactions, sufficient supply of nutrients, oxygen and growth factors, and no or minimal hydrodynamic stresses [9–13]. The current hPSC culturing methods, however, provide culturing conditions that are very different from these physiological microenvironments, leading to low culture efficiency and difficulty to culture cells at large scales. Two-dimensional (2D) cell culturing (e.g. with cell culture flasks) is the most widely used method for culturing hPSCs. However, a 2D environment is very different from an in vivo environment. Additionally, 2D culturing is labor-, space- and reagent-consuming, and considered only suitable for preparing small scale cells [1, 7, 8]. Three-dimensional (3D) suspension culturing (e.g. with stirred-tank bioreactors), in which cells are suspended in the culture medium, has been widely studied to scale up the production and has achieved some attractive results [1, 7, 8]. However, a critical problem with 3D suspension culturing is the uncontrolled cell agglomeration (figures S1(A) and (B) are available online at stacks.iop.org/BF/10/025006/mmedia) [1]. hPSCs have strong cell–cell interactions that make them aggregate [12, 13]. Suspended single hPSCs first associate to form small cell clusters (i.e., initial cell clustering), which is important for the survival and growth of dissociated hPSCs [12, 13]. These small cell clusters grow as spherical aggregates (i.e., spheroids) when cells divide. Spheroids, however, frequently fuse to each other to form large cell agglomerates (i.e., agglomeration) (figures S1(A) and (B)). Agglomeration leads to inhomogeneity in cell aggregate size and is detrimental to cell culture [1]. For instance, the transport of nutrients, oxygen, and growth factors to, and the metabolic waste from cells located at the center of large cell agglomerates (e.g., >400 μm diameter) become insufficient, leading to slow cell growth, apoptosis, and uncontrolled differentiation [1, 14]. While agitating the culture can enhance mass transport and reduce cell agglomeration, agitation also generates critical hydrodynamic stresses (e.g. shear stresses) that negatively impacts cells [1, 15, 16]. With the uncontrolled cell aggregation and hydrodynamic stresses, significant cell death, low cell growth and low volumetric yield are common in 3D suspension culturing [7]. For instance, we and others showed hPSCs typically expanded 4-fold in 4 days to yield around 2.0 × 10⁶ cells ml⁻¹ (table S1) [17–19]. These cells occupy ∼0.4% of the bioreactor volume [20].

Furthermore, the hydrodynamic conditions (e.g. the medium flow direction, velocity, shear force and chemical environment) generated by the agitation in 3D suspension culturing are very complicate [1, 7, 8, 15–19, 21]. They are spatially and temporally varied, and sensitive to many factors including the bioreactor design (e.g. impeller geometry, size and position, vessel geometry and size, positions of probes for pH, temperature, oxygen), medium viscosity and agitation rate [1, 21]. They are currently not well understood and hard to control [1, 15, 16, 21]. Additionally, how cells respond to these complicate hydrodynamic conditions is not well known and is hard to study [1, 15, 16]. These complicate conditions and knowledge gap lead to culture inconsistency and difficulty in scaling up, as shown by recent studies on producing cardiomyocytes from hPSCs in stirred-tank bioreactors [22, 23]. For three batches (∼100 ml suspension cultures) using a hESC line, the final yield ranged from 40 million to 100 million cells, and final cardiomyocyte purity ranged from 54% to 84%. Using the same bioreactor and a different hPSC line, the final yield ranged from 89 million to 125 million cells, and final cardiomyocyte purity ranged from 28% to 88% [22, 23]. When the culture volume was scaled from ∼100 ml to ∼1000 ml, the yield and differentiation efficiency were altered. Re-optimizing bioreactor design, agitation rate and culture protocol was required [22, 23]. This indicates the challenge of further scaling up the culture volume (e.g. to hundreds liters) since optimizing multiple factors in large culture volume is costly. To our best knowledge, the largest 3D suspension culture volume demonstrated to date for hPSCs is less than 10 l [1, 24].

Researchers have tried to encapsulate and culture hPSCs in various hydrogel scaffolds with the goal of using scaffolds as physical barriers to isolate cells from agglomeration and hydrodynamic stresses [7, 17, 25–29]. We recently found hPSCs could be effectively cultured in a very soft and thermoreversible hydrogel matrix (Mebiol^® Gel) [7, 17, 29]. Within this hydrogel matrix, single hPSCs clonally grew into uniform spheroids (diameter ∼100–150 μm) (figures S1(C) and (D)). The hydrogel scaffold enabled serial expansion of hPSCs with good cell viability, growth rate (e.g. 20-fold/5 days), yield (e.g. 2.0 × 10⁷ cells ml⁻¹) and purity, all of which offered improvements over 3D suspension cultures. However, this system has some limitations that make it inappropriate for large-scale cell production. First, it is not universally applicable. For some hPSC lines and human cell types, the dissociated cells cannot survive and clonally grow in the hydrogel. Though the exact reason is still unknown, we speculate that the scaffold prevents the initial cell–cell interactions that are important for cell's survival [7, 17, 29]. Second, the volumetric yield and expansion per passage are moderate, probably due to the fact that cells encapsulated in hydrogels have no free space for expansion and they have to deform scaffolds to create spaces for the new cells [7, 17, 29]. Third, the hydrogel is mechanically fragile, making it hard to be processed into formats (e.g., beads) suitable for large-scale cell cultures. And lastly, the hydrogel remains stable for about two weeks, which prohibits its use for long-term cell culture, such as differentiating hPSCs into neurons, which requires more than one month. The results of this study showed that eliminating the cell agglomeration and hydrodynamic stresses could significantly improve the culture efficiency, however, free spaces that allow the initial cellular interactions and subsequent cell expansion were also important to achieve high culture outcomes.

We here report a novel technology that can overcome the limitations of current hPSC culturing methods and provide physiologically relevant cell culture microenvironments. With this technology, hPSCs are processed into and cultured in microscale alginate hydrogel tubes (or AlgTubes) that are suspended in the cell culture medium in a culture vessel (figures 1(A) and (B)). The hydrogel tubes create cell-friendly microspaces that allow cells to interact with each other and expand. Meanwhile, they protect cells from hydrodynamic stresses in the culture vessel and confine the cell mass less than 400 μm (in radial diameter) to ensure efficient mass transport during the entire culture (figures 1(A) and (B)). Additionally, this technology is simple, scalable, defined and compatible with the current good manufacturing practices that make it commercially viable. We showed that, under optimized culture conditions, the AlgTubes offered paradigm-shifting improvements in cell viability, growth, yield, culture consistency and scalability over current hPSC culturing technologies.

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Materials

Culturing hPSCs in 3D PNIPAAm-PEG hydrogel

Processing AlgTubes

Culturing hPSCs in the AlgTubes

Culturing L-Wnt3A-cells in AlgTubes

Culturing hPSCs in the AlgTubes with bioreactor

Staining, flow cytometry and imaging

EB differentiation

Teratoma formation in vivo

Karyotype

Mesodermal induction

Endodermal induction

Cardiomyocyte differentiation

Direct comparison of culturing hPSCs in 2D, static 3D and dynamic 3D suspension and the AlgTubes

RNA sequencing and data analysis

Principle component analysis (PCA) analysis

Statistical analysis

Data availability

A micro-extruder was designed and made for processing AlgTubes (figure 1). To make AlgTubes, a cell solution and an alginate solution is pumped into the central channel and side channel of the micro-extruder, respectively, to form coaxial core–shell flows that are extruded into a CaCl₂ buffer. The shell alginate flow is instantly crosslinked by Ca²⁺ ions to form an alginate hydrogel tube (figures 1(C)–(E)). Subsequently, the CaCl₂ buffer is replaced by the cell culture medium and cells are grown in the tubes. The micro-extruder can have multiple nozzles for simultaneously processing multiple AlgTubes to make large-scale AlgTubes (figure S2, and supplementary video 1). It was found that the cell solution and alginate solution should have close viscosity to process defect-free AlgTubes. Both hyaluronic acid (HA) and methylcellulose (MC) solutions could be used to suspend cells for this purpose. Similar to 3D suspension culturing (figures S1(A) and (B)), single hPSCs in AlgTubes quickly (e.g. within 24 h) associated to form small cell clusters that subsequently grew and filled the tubes (figures 1(F) and (G)). We termed these two phases as the initial clustering phase and subsequent cell expansion phase, respectively. In 2D and 3D suspension culturing of hPSCs, ROCK inhibitors (e.g. Y-27632) are added to the culture medium for the first 24 h after passaging to inhibit the dissociation-induced cell death [30]. We found ROCK inhibitors (Y-27632) were also required for the survival and growth of the single hPSCs in AlgTubes. Without ROCK inhibitors, majority of cells died within 24 h (figures 1(H) and (I)). To passage cells, the AlgTubes could be dissolved with the cell-compatible ethylenediaminetetraacetic acid (EDTA) solution (0.5 mM, 5 min at room temperature) to release the micro cell mass, which could be further treated with Accutase (10 min at 37 °C) and dissociated into single cells for the following passage.

We found AlgTubes made with a wide range of alginate concentrations, tube diameters and hydrogel shell thicknesses all supported efficient culturing of hPSCs (figures S3–S5). For instance, hPSCs grew efficiently and similarly in AlgTubes made with 1.0%, 1.5% and 2.0% alginate solutions (figure S3). When seeded at 1.0 × 10⁷ cells ml⁻¹, cells expanded ∼14-, 23- and 50-fold to yield ∼1.4×, 2.3 × and 5.0 × 10⁸ cells ml⁻¹ on day 5, 7 and 9 of the culturing with above 94% cells expressed the pluripotency marker Oct4 at all three alginate concentrations. The AlgTube's outer diameter is roughly equal to the inner diameter of the nozzle of the micro-extruder and can be controlled through adjusting the nozzle's diameter (figure S2). At a given AlgTube outer diameter, the hydrogel shell thickness could be controlled by varying the ratio of the cell solution flow rate and alginate solution flow rate and predicted by using the equation described in figure S4(A). When seeded at 1.0 × 10⁷ cells ml⁻¹ and cultured in AlgTubes (with ∼400 μm diameter) with 30, 40, and 70 μm shells or AlgTubes (with 30–50 μm shells) with diameter of 400 μm, 250 μm and 120 μm for 9 days, hPSCs all expanded around 50-fold to yield ∼5.0 × 10⁸ cells ml⁻¹ with above 90% cells retained the pluripotency marker Oct4 (figures S4 and S5). Since both HAs and MCs could be used to suspend cells, we studied whether they differentially influenced cell culture. Our research showed HAs and MCs resulted in similar cell viability, expansion and pluripotency (figure S6). The capability to support consistent hPSC culturing under a wide range of AlgTube conditions gives users large flexibility in processing the tubes.

AlgTubes supported efficient hPSC culturing with a wide range of seeding densities (figures 2 and S7). When seeded at 1.0×, 2.0×, 5.0×, and 10.0 × 10⁶ cells ml⁻¹, hPSCs expanded 499-, 247-, 104- and 51-fold by day 9, respectively, yielding around 5.0 × 10⁸ cells ml⁻¹ (figure 2). For all conditions, cells grew through the initial clustering phase and the subsequent cell expansion phase. At 24 h after processing the AlgTubes, the cell cluster size was larger for higher seeding density but the numbers of cell clusters per volume were similar for different seeding densities (figure 2(A)). These results showed hPSCs grew faster at lower seeding density (figures 2(B) and (C)). However, the seeding density did not influence pluripotency. >95% cells expressed the pluripotency marker Oct4 for all seeding densities (figures 2(D) and (E)). It is extremely exciting that hPSCs seeded at ultralow densities could grow as well without sacrificing cell viability and pluripotency (figure S7). When seeded at 1.0×, 3.0 × and 5.0 × 10⁵ cells ml⁻¹, hPSCs expanded 4200-, 1700- and 1000-fold to yield ∼4.2×, 5.1×, and 5.0 × 10⁸ cells ml⁻¹ by days 14, 12 and 10, respectively.

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After performing these optimizations, the AlgTubes were evaluated for culturing multiple hPSC lines for long-term (e.g. 10 passages, figures 3, S8 and S9). All hPSCs grew well in AlgTubes, and cell morphology, viability, growth rate, and pluripotency among multiple hPSC lines were similar. In the 10-passage culture, when seeded at 1.0 × 10⁷ cells ml⁻¹, hPSCs consistently expanded ∼15-fold/passage/5 days and >95% of cells expressed the pluripotency marker Oct4 (figures 3(A) and (B)). The long-term culture did not alter cell phenotype as shown by the similar cell morphology, viability, growth kinetics and pluripotency of hPSCs at passage 1 and 10 (figures 3, S8 and S9). When seeded at 1.0 × 10⁷ cells ml⁻¹, hPSCs expanded ∼50-fold to yield 5.0 × 10⁸ cells ml⁻¹ on day 9 at both passage 1 and 10. In vitro embryoid body (EB) differentiation and in vivo teratoma formation confirmed their pluripotency after the long-term culture. All hPSCs were successfully differentiated into endodermal, mesodermal and ectodermal cells in the EB assay (figures 4(A), S10(A) and (B)). All hPSCs formed teratomas containing the three germ layer tissues in immune-deficient mice in the teratoma assay (figures 4(B), S10(C) and (D)). In addition, after long-term culture, all hPSCs retained normal karyotypes (figures 4(C), S10(E) and (F)). These results show that AlgTubes can support long-term culturing of hPSCs.

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After expansion and further culturing in a mesodermal [31, 32] or endodermal [33] or cardiomyocyte [31, 32] differentiation medium, hPSCs in the AlgTubes could be differentiated to corresponding mesodermal or endodermal cells or cardiomyocytes at high efficiency, indicating the AlgTubes support hPSC differentiation (figures 4(D)–(F), and supplementary video 2). Additionally, cells other than hPSCs can be cultured in the AlgTubes. For instance, murine L cells engineered to express Wnt3A proteins were efficiently cultured without notable cell death [34], yielding around 6.0 × 10⁸ cells ml⁻¹ (figure S11). Wnt3A proteins were consistently expressed during a 16 day culture. These results demonstrate AlgTube' potential as a generally applicable culture system.

A prototype bioreactor was built to show AlgTubes' scalability (figure 5). Two milliliters of AlgTubes with hPSCs were processed and contained in a bioreactor. The cell culture medium was stored in a plastic bellow bottle that could be pressed to flow the medium into the bioreactor or released to withdraw the medium from the bioreactor, respectively (figure 5(A)). The pressing and releasing speed, as well as the duration of the interval between the pressing and releasing can be programmed and controlled by the controller (figure 5(B)). Since AlgTubes have similar density with the cell culture medium, they were uniformly suspended and dispersed in the medium when the medium was pumped into the bioreactor. They became collected and contact with each other when the medium was withdrawn from the bioreactor. This alternating dispersion and collection is designed to enhance the medium mixing. Within the bioreactor, hPSCs grew well and yielded ∼5.0 × 10⁸ cell ml⁻¹ by day 10 (figures 5(C) and (D)). >95% of cells expressed the pluripotency markers (figures 5(E) and (F)). This prototype bioreactor could be further scaled up in the future.

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To assess the superiority of the AlgTubes over current hPSC culturing technologies, we directly compared culturing hPSCs in 2D, 3D suspension without agitation (static 3D), 3D suspension with agitation (dynamic 3D) and the AlgTubes (figure 6). Cells were cultured for 9 days in AlgTubes to allow cells to fill the tubes. Cells were cultured for 7 days in 2D, static 3D and dynamic 3D culturing since the cell numbers had no further increase after 5 days in these cultures. Medium was changed daily. Live cells in the cultures were counted daily. Cells died in each day were quantified through measuring the adenylate kinases (AKs) in the exhausted cell culture medium. AKs, which are ubiquitous proteins presented in cells, are rapidly released into the culture medium upon damage of the plasma membrane. The ratio of dead cells over the dead cells plus the live cells was calculated (e.g. the % dead cells) and used to assess the cell viability each day. Cell proliferation was also evaluated through quantifying the percentage of cells in S and G2/M phases. Seeded at 2.0 × 10⁵ cells ml⁻¹, hPSCs expanded only 5-fold to yield a maximum of ∼1.0 × 10⁶ cells ml⁻¹ in static 3D culturing. At the same seeding density, hPSCs expanded ∼9-fold to yield a maximum of ∼1.8 × 10⁶ cells ml⁻¹ in dynamic 3D culturing. In 2D, cells could expand ∼51-fold to generate ∼1.0 × 10⁷ cells per well of the 6-well plate. In AlgTubes, hPSCs expanded ∼500-fold to yield ∼5.0 × 10⁸ cells ml⁻¹ (figures 6(A) and (B)). Large cell death was observed in static 3D, dynamic 3D and 2D culturing, especially after day 3 or when the cell density reached ∼1.0 × 10⁶ cells ml⁻¹ (figures 6(D) and (E)). In static 3D culturing, the % of dead cells reached ∼43%, 53%, 59% and 62% on day 4, 5, 6 and 7, respectively. In dynamic 3D culturing, the % of dead cells reached ∼28%, 29%, 35% and 40% on day 4, 5, 6 and 7, respectively. In AlgTubes, cell death was low and the % of dead cells was <10% during the entire 9 day culture. Cell cycle analysis showed static 3D, dynamic 3D and 2D culturing did not decrease the % of cells in proliferation (figure 6(C)). To study whether the global gene expression are different, mRNAs of the day 3 cells from the 4 culturing methods were sequenced. On day 3, cell death was not significant and majority of cells were proliferating in the 4 cultures (figures 6(C)–(E)). Hierarchical clustering analysis showed the gene expression profiles were very similar between the 4 cultures (figures 6(F) and S12). These results show that large cell death in 3D suspension culturing leads to low culture efficiency, and the AlgTubes improve the culture outcome through reducing the cell death without significantly altering the global gene expression and cell phenotype.

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We also directly compared encapsulating and culturing hPSCs in solid agarose, HA, and alginate hydrogels with suspending and culturing hPSCs in AlgTubes (figure S13). When seeded at 1 × 10⁶ cells ml⁻¹, little cell expansions were observed in HA and agarose hydrogels. Small numbers of spheroids were formed and hPSCs expanded ∼3.5-fold in solid alginate hydrogel fibers. In AlgTubes, hPSCs expanded ∼500-fold to yield ∼5.0 × 10⁸ cells ml⁻¹. These results show the importance of the free microspaces for achieving high cell expansion and yield.

Based on the results of culturing hPSCs in 2D, 3D suspension and hydrogels [7, 17, 25–29], we proposed that 3D culture microenvironments having free space for the initial cell–cell interactions and subsequent cell growth, but no cell agglomeration and hydrodynamic stresses could significantly improve the culture efficiency. The AlgTubes are designed to provide these physiologically relevant microenvironments (figures 1(A) and (B)). First, cells in AlgTubes are protected from hydrodynamic stresses/conditions by the hydrogel shells. This reduces the hydrodynamic-conditions-induced negative effects [1, 15, 16, 21–23]. Second, cell masses in AlgTubes are controlled to be less than 400 μm to ensure efficient mass transport during the entire culture. Additionally, the cell masses are monodisperse (in radial diameter), which can improve the homogeneity of hPSC expansion and differentiation. Other methods, such as culturing cells with AggreWell^™, can precisely control cell aggregate size, however, are not scalable. Third, unlike encapsulating and culturing cells in hydrogels (figure S13), the AlgTubes provides free space for hPSC expansion and does not hinder the initial cellular interaction, leading to high volumetric yield and general applicability.

The use of alginates makes this new technology scalable, cost-effective and GMP-compatible. Alginates are affordable; available in large quantity; non-toxic to cells; have been used in clinics; and can be instantly crosslinked with Ca²⁺ ions to process large-scale AlgTubes with the scalable extrusion technology (figure S2) [35]. The resulting AlgTubes are mechanically and chemically stable for months and suitable for large-scale and long-term cell cultures; can be dissolved easily with cell-compatible EDTA solution to release the product; and are transparent to allow observation of cell growth [36].

The conceptual and technical innovations of AlgTubes lead to high culture efficiency. We demonstrated long-term culturing (>10 passages) of multiple hPSC lines without uncontrolled differentiation and chromosomal abnormalities (figures 3, S8 and S9). Cultures between batches and cell lines were very consistent (figure 3(A)). hPSCs in AlgTubes had high viability, growth rate (1000-fold/10 days/passage in general) and yield (∼5 × 10⁸ cells ml⁻¹ microspace). The expansion per passage (e.g. up to 4200-fold/passage was achieved in figure S7) and volumetric yield are much higher than current 3D suspension culturing (table S1) [7, 8, 17, 37, 38]. For comparison, in 3D suspension culturing, we typically achieve 4-fold expansion per 4 days to yield around 2.0 × 10⁶ cells ml⁻¹. As discussed later, the high yield and high expansion fold have high impact on large-scale cell production since they significantly reduce the culture volume and time, numbers of passaging operations, and the production cost. hPSCs could be efficiently differentiated into various tissues cells (figure 4 and supplementary video 2). In addition, AlgTubes-based scalable bioreactors could be readily built (figure 5). Our comparative study showed the AlgTubes did not significantly alter hPSCs' gene expression profiles, but significantly reduced cell death, resulting in high cell expansion and yield (figures 6 and S12).

It should be noted that pluripotent stem cells have been cultured in hollow fibers made from dry polymer membranes [39–41]. However, their performance offers no advancements over stirred-tank bioreactors. In addition, harvesting cells from these fibers were very challenging. For instance, it required treating the cells with trypsin for 30–60 min. Moreover, cells could not be observed with microscopy and sampling was hard during the culture. Solid hydrogel fibers (e.g. not hollow) have been studied for processing tissues [42–44]. However, the AlgTubes are hollow and conceptually and technically different from these systems.

The AlgTubes will be of broad interest to individual laboratories, institutions, and biotechnology companies working on regenerative medicine and cancer therapies. First, AlgTubes provide laboratories an easy-to-use tool for studying biology in 3D, which can better replicate the in vivo biology than 2D cultures [45]. For instance, AlgTubes may be used to culture organoids [46, 47], study cellular assembly into tissues [48] and reprogram cells [49], etc. Second, the AlgTubes can greatly benefit translational research. Using hPSC-derived cardiomyocytes for treating myocardial infarction as an example [50], 2D culturing was first used for developing the differentiation protocol and the initial demonstration of efficacy and safety in rodents [51]. Subsequently, large effort was spent to develop 3D suspension culturing for preparing sufficient cardiomyocytes for tests on large animals [52, 53]. If successful, substantial amounts of effort will be required to develop a new and large bioprocess for making sufficient cells for clinical trials and eventually clinics [50]. Since cell's phenotype, safety and efficacy are bioprocess-related, extensive and expensive in vitro and in vivo characterization of cell phenotype, safety, and efficacy are required after each change of the culture method [54]. These efforts make developing cellular therapeutics slow and expensive. AlgTubes is scalable and can be used for culturing cells for all these development stages. For instance, ∼200 μl AlgTubes can be used for developing the differentiation protocol. ∼2 ml AlgTubes are sufficient to produce ∼10⁹ cells for preclinical small animal studies. ∼200 ml AlgTubes can yield ∼10¹¹ cells for tests on large animals and clinical trials. The AlgTubes can save significant time and investment for developing cellular therapeutics. Third, the AlgTubes are very attractive for industry-scale cell production. A simple comparative calculation of producing ∼1.5 × 10¹⁴ hPSCs (from ∼10⁸ hPSC seeds) with stirred-tank bioreactors and AlgTubes shows the significant impact of AlgTubes' high cell expansion per passage and volumetric yield (figure S14). For the calculation, we assumed 4-fold expansion per 4 days per passage for stirred-tank bioreactors and 1000-fold expansion per 10 days per passage for AlgTubes as well as a seeding density of 5 × 10⁵ cells ml⁻¹ and a passaging efficiency (i.e., % of cells remaining viable after one passaging) of 80% for both. These assumptions are based on data from my lab and others (table S1) [7, 17, 24, 55]. The production requires ∼104, 811 liters of total culture volume, 11 passaging operations, and 48 days with stirred-tank bioreactors, which is technically and economically challenging (figures S14(A) and (C)). As mentioned, building large stirred-tank bioreactors will be very challenging. Passaging is highly unwanted for large-scale cell production since it requires large amounts of time, labor, and reagent; increases the risk of contamination and culture failure; loses large numbers of cells; and causes difficulties during automation. The production can be done with 320 l of AlgTubes in 20 days and 1 passaging (figures S14(B) and (C)). The reductions in culture volume, time, and passaging not only make the production technically feasible, but also lead to enormous cuts in labor, reagents, equipment, cGMP facility size, and overall production cost.

In summary, the AlgTube technology combines physiologically relevant culture microenvironments, high performance, high scalability, cGMP compliance and commercial viability, and has potential to address the hPSC manufacturing challenge. Advances in biology have developed protocols for efficiently differentiating hPSCs into many human cell types [7, 31]. Future research can explore integrating these protocols into AlgTubes to produce various human cell types. It will be valuable to systematically study culturing other cell types, such as human adult stem cells and T cells in AlgTubes.

Aaron Howell at the Department of Chemical and Biomolecular Engineering at University of Nebraska, Lincoln drew the 3D graphics. Leonard Akert at the Department of Chemical and Biomolecular Engineering at University of Nebraska, Lincoln assisted QL with building the extruders and prototype bioreactors. Drs Stephen D Kachman and Zheng Xu at the Department of Statistics, University of Nebraska, Lincoln assisted the statistical analysis. Confocal microscope imaging was done in the Morrison Microscopy Core Research Facility at University of Nebraska, Lincoln. Drs You Zhou and Christian Elowsky assisted the confocal imaging. Flow cytometry was done in the Morrison center, the Flow Cytometry core, University of Nebraska, Lincoln with the assistance of Dirk Anderson. RNA sequencing was done at the UNMC deep sequencing core. Drs Hendrik J Viljoen and Anuradha Subramanian at the Department of Chemical and Biomolecular Engineering at University of Nebraska, Lincoln offered discussion and advice on the manuscript.

YL conceived the idea. YL and CZ directed the work. YL and QL designed the experiments and wrote the manuscript. QL, HL, OW, CE and HC performed experiments. QL analyzed the data. QD, KL and CZ analyzed the sequencing data.

The University of Nebraska has filed a patent application related to the technology described in this work to the United States Patent and Trademark Office.