Enhanced oxygenation of insulin-secreting cells by a bioelectronic-assisted macroencapsulation device (beamed)

Abstract

Disclosed herein are an implantable therapeutic delivery system, methods of making the same, and methods of using the same. This system includes an electrochemical oxygen generator and a cell compartment comprising a semi-permeable nanofibrous membrane that is impermeable to cellular migration, an internal hydrogel matrix and one or more cells embedded in the hydrogel matrix. This system further includes a conduit in fluid communication between the electrochemical oxygen generator and the interior of the cell compartment, wherein the conduit delivers oxygen generated by the electrochemical oxygen generator to the one or more cells embedded in the hydrogel matrix.

Description

ENHANCED OXYGENATION OF INSULIN-SECRETING CELLS BY A BIOELECTRONIC-ASSISTED MACROENCAPSULATION DEVICE (BEAMED)

[0001] This application claims the priority benefit of U.S. Provisional Patent Application Serial No.63 / 702,831, filed October 3, 2024, which is hereby incorporated by reference in its entirety. FIELD OF THE INVENTION

[0002] The present invention relates to methods and devices for promoting the enhanced oxygenation of insulin-secreting cells by a bioelectronic-assisted macroencapsulation device (BEAMED). BACKGROUND

[0003] Type 1 diabetes (T1D) is a chronic disorder caused by the autoimmune destruction of pancreatic β-cells, which eventually leads to insulin deficiency. Exogenous insulin therapy, the current mainstay to manage this disease, necessitates a stringent and constant effort from patients to carefully manage insulin intake balanced with diet, exercise, sleep, and stress. Despite the significant advances in insulin therapy, it remains non-curative and incapable of maintaining glucose homeostasis, predisposing patients to life-threatening complications including severe hypoglycemia events (SHEs) (Dagogo-Jack, S., “Hypoglycemia in Type 1 Diabetes Mellitus: Pathophysiology and Prevention,” Treat Endocrinol 3(2):91-103 (2004); Pedersen-Bjergaard et al., “Severe Hypoglycaemia in 1076 Adult Patients with Type 1 Diabetes: Influence of Risk Markers and Selection,” Diabetes Metab Res. Rev.20(6):479-86 (2004); Cooper et al., “Clinical and Demographic Risk Factors Associated with Mortality During Early Adulthood in a Population-Based Cohort of Childhood-Onset Type 1 Diabetes,” Diabet. Med.31(12):1550-8 (2014)) and irreversible organ damage caused by chronic hyperglycemia (Desai et al., “Advances in Islet Encapsulation Technologies,” Nature Reviews Drug Discovery 16(5):338-350 (2017)). Cellular therapy has emerged as a potential cure to overcome this challenge given the ability of primary and stem cell-derived insulin-producing cells to inherently respond to blood glucose changes. Multicenter clinical trials of intraportal transplantation of allogenic cadaveric islets in over 1200 patients have consistently demonstrated the superior efficacy and safety of cell therapy, evidenced by physiological glycemic control without exogenous insulin and nearly complete eradication of SHEs (Chetboun et al., “Association Between Primary Graft Function and 5-Year Outcomes of Islet Allogeneic Transplantation in Type 1 Diabetes: A Retrospective, Multicentre, Observational Cohort Study in 1210 Patients from The Collaborative Islet 320349050v3Transplant Registry,” The Lancet Diabetes & Endocrinology 11(6):391-401 (2023); Hering et al., “Factors Associated with Favourable 5 Year Outcomes in Islet Transplant Alone Recipients with Type 1 Diabetes Complicated by Severe Hypoglycaemia in the Collaborative Islet Transplant Registry,” Diabetologia 66(1):163-173 (2023)). Favorable outcomes of the recent Phase 1 / 2 clinical trial by Vertex Pharmaceuticals (NCT04786262) further confirmed the feasibility of cell replacement therapy for T1D with the limitation of donor availability being addressed by the sustainable supply of stem cell-derived β cells (SC-β cells). However, the requisite lifelong immunosuppression with deleterious side effects, such as nephrotoxicity, opportunistic infections, and malignancies, remains a formidable obstacle to the widespread clinical application of current cellular therapies (Ryan et al., “Risks and Side Effects of Islet Transplantation,” Curr. Diab. Rep.4(4):304-9 (2004)). Thus, developing an immunosuppression-free strategy to protect the transplanted cells is imperative to fully harness the potential of cell therapy for curing T1D.

[0004] Over the past few decades, the concept of cell encapsulation has been proposed to circumvent immunosuppression. In principle, enveloping insulin-secreting cells within a semipermeable system could prevent direct contact with the detrimental elements of the immune system while allowing for efficient diffusion of glucose, insulin, nutrients, and metabolic wastes. Among encapsulation strategies, macroencapsulation recently garnered more attraction in the field, especially after SC-β cells became clinically available. As opposed to microencapsulation, cell confinement and the ability to retrieve the whole system of the macroencapsulation devices can address safety and regulatory concerns. Furthermore, subcutaneous space stands out as an ideal transplantation site given the large capacity, minimally invasive surgical procedure, amenability to frequent monitoring, and safe retrievability. However, despite notable insights from research endeavors, translating this concept into clinical applications has been impeded by persistent and intertwined challenges including insufficient supply of oxygen, foreign body responses (FBR), and inability of maintaining a curative dose of cells functional in a reasonably sized system. Particularly, β-cells have an intrinsically high oxygen consumption rate due to the high metabolic demand for insulin synthesis and secretion. Native pancreatic islets require a disproportionately high proportion of pancreatic blood flow relative to their mass (Colton, C.K., “Oxygen Supply to Encapsulated Therapeutic Cells,” Advanced Drug Delivery Reviews 67- 68:93-110 (2014); Carlsson et al., “Markedly Decreased Oxygen Tension in Transplanted Rat Pancreatic Islets Irrespective of the Implantation Site,” Diabetes 50(3):489-95 (2001)). However, the oxygen supply in encapsulation systems is inadequate due to the lack of direct vascularization of encapsulated cells and relatively low oxygen tension in extrahepatic sites 320349050v3(such as the desirable subcutaneous milieu), leading to hypoxia-induced cell death (Avgoustiniatos, E.S., “Oxygen Diffusion Limitations in Pancreatic Islet Culture and Immunoisolation,” Thesis, Massachusetts Institute of Technology (2002); Avgoustiniatos et al., “Measurements of the Effective Diffusion Coefficient of Oxygen in Pancreatic Islets,” Industrial & Engineering Chemistry Research 46(19):6157-6163 (2007); Suzanne, L.A., “Eliminating Oxygen Supply Limitations for Transplanted Microencapsulated Islets in the Treatment of Type 1 Diabetes,” Thesis, Massachusetts Institute of Technology (2008)) and irreversibly impaired β- cell function (Dionne et al., “Effect of Hypoxia on Insulin Secretion by Isolated Rat and Canine Islets of Langerhans,” Diabetes 42(1):12-21 (1993); Cantley et al., “A Preexistent Hypoxic Gene Signature Predicts Impaired Islet Graft Function and Glucose Homeostasis,” Cell Transplant 22(11):2147-59 (2013)). The oxygen deficit is further exacerbated by the low oxygen permeability of fibrotic tissues surrounding the implants caused by foreign body response against the encapsulating materials (Kumosa et al., “Permeability of Subcutaneous Tissues Surrounding Long-Term Implants to Oxygen,” Biomaterials 35(29):8287-8296 (2014)). Moreover, insufficient oxygen supply also limits the packing density of β-cells within the encapsulation system as escalating cell density amplifies the requirement for oxygen owing to the steeper oxygen gradient among cell layers (Pedraza et al., “Preventing Hypoxia-Induced Cell Death in Beta Cells and Islets via Hydrolytically Activated, Oxygen-Generating Biomaterials,” Proc. Natl. Acad. Sci. U.S.A.109(11):4245-50 (2012); Coronel et al., “Mitigating Hypoxic Stress on Pancreatic Islets via in Situ Oxygen Generating Biomaterial,” Biomaterials 129:139-151 (2017); Coronel et al., “Oxygen Generating Biomaterial Improves the Function and Efficacy of Beta Cells within a Macroencapsulation Device,” Biomaterials 210:1-11 (2019); Gholipourmalekabadi et al., “Oxygen-Generating Biomaterials: A New, Viable Paradigm for Tissue Engineering?” Trends in Biotechnology 34(12):1010-1021 (2016); Farris et al., “Oxygen Delivering Biomaterials for Tissue Engineering,” J Mater Chem B 4(20):3422-3432 (2016)). As a result, low cell densities are typically used to reduce hypoxia-induced cell loss post- implantation, rendering the requisite implant dimensions impractical for delivering a curative dose of β-cells (Colton, C.K., “Oxygen Supply to Encapsulated Therapeutic Cells,” Advanced Drug Delivery Reviews 67-68:93-110 (2014); de Vos et al., “Encapsulation of Pancreatic Islets for Transplantation in Diabetes: The Untouchable Islets,” Trends in Molecular Medicine 8(8):363-366 (2002)). This limitation in packing density has not always been noted in small animal studies but poses a significant barrier to scaling up for clinical translation.

[0005] Enhanced oxygen supply to cell encapsulation systems could potentially address the aforementioned challenges. Integration of oxygen-generating biomaterials (e.g. calcium peroxide 320349050v3(Pedraza et al., “Preventing Hypoxia-Induced Cell Death in Beta Cells and Islets via Hydrolytically Activated, Oxygen-Generating Biomaterials,” Proc. Natl. Acad. Sci. U.S.A. 109(11):4245-50 (2012); Coronel et al., “Oxygen Generating Biomaterial Improves the Function and Efficacy of Beta Cells within a Macroencapsulation Device,” Biomaterials 210:1-11 (2019); Liang et al., “Engineering a Macroporous Oxygen-Generating Scaffold for Enhancing Islet Cell Transplantation within an Extrahepatic Site,” Acta Biomater 130:268-280 (2021)), sodium percarbonate (Ward et al., “Oxygen Generating Biomaterials Preserve Skeletal Muscle Homeostasis Under Hypoxic and Ischemic Conditions,” PLoS One 8(8):e72485 (2013); Harrison et al., “Oxygen Producing Biomaterials for Tissue Regeneration,” Biomaterials 28(31):4628-34 (2007)), and lithium peroxide (Wang et al., “An Inverse-Breathing Encapsulation System for Cell Delivery,” Science Advances 7(20):eabd5835 (2021)) into the encapsulation systems has shown encouraging outcomes in preclinical models. The common downsides of these chemical- based approaches are the short duration of oxygen supply, uncontrolled oxygen generation rate, and undesirable end products (Pedraza et al., “Preventing Hypoxia-Induced Cell Death in Beta Cells and Islets via Hydrolytically Activated, Oxygen-Generating Biomaterials,” Proc. Natl. Acad. Sci. U.S.A.109(11):4245-50 (2012); Gholipourmalekabadi et al., “Oxygen-Generating Biomaterials: A New, Viable Paradigm for Tissue Engineering?” Trends in Biotechnology 34(12):1010-1021 (2016)). Another approach, by BetaO2 Technologies, tackled these challenges by integrating a refillable oxygen reservoir into a macroencapsulation system, allowing for oxygen replenishment via percutaneous injection ports. Devices supplied daily with supraphysiological levels of oxygen demonstrated long-term diabetes reversal in both an allogeneic rat-to-rat and a xenogeneic rat-to-pig model (Colton, C.K., “Implantable Biohybrid Artificial Organs,” Cell Transplant 4(4):415-36 (1995); Avgoustiniatos et al., “Effect of External Oxygen Mass Transfer Resistances on Viability of Immunoisolated Tissue,” Ann NY Acad Sci 831:145-67 (1997); Papas et al., “Oxygenation Strategies for Encapsulated Islet and Beta Cell Transplants,” Advanced Drug Delivery Reviews 139:139-156 (2019)). Nonetheless, the requirement for oxygen replenishing via frequent injections and device bulkiness pose ongoing challenges.

[0006] The present invention is directed to overcoming these and other deficiencies in the art. SUMMARY

[0007] A first aspect of the present disclosure relates to an implantable therapeutic delivery system. This system comprises an electrochemical oxygen generator; a cell compartment 320349050v3comprising a semi-permeable nanofibrous membrane that is impermeable to cellular migration, an internal hydrogel matrix and one or more cells embedded in the hydrogel matrix; and a conduit in fluid communication between the electrochemical oxygen generator and the interior of the cell compartment, wherein the conduit delivers oxygen generated by the electrochemical oxygen generator to the one or more cells embedded in the hydrogel matrix.

[0008] A second aspect of the present disclosure relates to a method of delivering a therapeutic agent to a subject in need thereof. This method involves implanting the implantable therapeutic delivery system according to the first aspect into the subject.

[0009] A third aspect of the present disclosure relates to a method of treating diabetes in a subject. This method involves implanting the implantable therapeutic delivery system according to the first aspect into the subject having diabetes.

[0010] A fourth aspect of the present disclosure relates to a method of treating a bleeding disorder in a subject. This method involves implanting the implantable therapeutic delivery system according to the first aspect into the subject having a bleeding disorder.

[0011] A fifth aspect of the present disclosure relates to a method of treating a lysosomal storage disease in a subject. This method involves implanting the implantable therapeutic delivery system according to the first aspect into the subject having the lysosomal storage disease.

[0012] A sixth aspect of the present disclosure relates to a method of treating a neurological disorder in a subject. This method involves implanting the implantable therapeutic delivery system according to the first aspect into the subject having the neurological disorder.

[0013] A seventh aspect of the present disclosure relates to a method of treating a cancer in a subject. This method involves implanting the implantable therapeutic delivery system according to the first aspect into the subject having cancer.

[0014] An eighth aspect of the present disclosure relates to a method of treating a chronic eye disease in a subject. This method involves implanting the implantable therapeutic delivery system according to the first aspect into the subject having a chronic eye disease.

[0015] A ninth aspect of the present disclosure relates to a method of treating a kidney failure in a subject. This method involves implanting the implantable therapeutic delivery system according to the first aspect into the subject having a kidney failure.

[0016] A tenth aspect of the present disclosure relates to a method of treating a chronic pain in a subject. This method involves implanting the implantable therapeutic delivery system according to the first aspect into the subject having a chronic pain. 320349050v3

[0017] An eleventh aspect of the present disclosure relates to a method of forming a seamless nanofibrous membrane. This method involves forming an enlarged diameter support about a mandrel, the support being formed of a dissolvable material; and electrospinning a nanofibrous membrane about the enlarged diameter support and a portion of the mandrel.

[0018] A twelfth aspect of the present disclosure relates to a method of making a cell encapsulation system suitable for implant. This method involves providing a semi-permeable nanofibrous membrane structure; inserting a permeable conduit into the semi-permeable nanofibrous membrane, the permeable conduit having an unsealed end; and introducing cells and hydrogel precursor via the permeable conduit, whereby the cells and hydrogel precursor are forced into the space between the permeable conduit and the nanofibrous membrane, followed by crosslinking of the hydrogel precursor to form the hydrogel matrix. This method further involves joining together the unsealed end of the permeable conduit to a distal end of an impermeable conduit coupled to an electrochemical oxygen generator; and sealing the semi- permeable nanofibrous membrane about the distal end of the impermeable conduit.

[0019] A thirteenth aspect of the present disclosure relates to a device for forming an implantable therapeutic delivery system. This device comprises an electrochemical oxygen generator; a compartment comprising a semi-permeable nanofibrous membrane that is impermeable to cellular migration; a conduit in fluid communication between the electrochemical oxygen generator and the interior of the compartment, wherein the conduit delivers oxygen generated by the electrochemical oxygen generator to the interior of the compartment.

[0020] A fourteenth aspect of the present disclosure relates to a kit. This kit comprises the device according to the thirteenth aspect; one or more precursors for forming a hydrogel matrix; and instructions for loading a composition comprising cells and the one or more precursors into the compartment of the device, and for forming the hydrogel matrix in situ.

[0021] To address the problems associated with prior cell encapsulation systems, a BioElectronics-Assisted Macroencapsulation (BEAM) system capable of providing a continuous oxygen supply to encapsulated cells via electrolysis of tissue moisture was developed. The components of the implantable electrochemical oxygen generator (iEOG) are assembled in a compact, enclosed design (13 mm diameter x 3.1 mm thick), which is then connected to a linear core-shell cell pouch. Islets are housed within the annular space between an inner gas-permeable silicone tubing and an outer immunoprotective tubing made of a hydrogel-impregnated electrospun membrane. This unique design allows for optimal mass transportation and proximity between cells and the oxygen source. The concentric configuration augments the surface-to- 320349050v3volume ratio and facilitates scalability both longitudinally and radially. The edge-free, round-end cylindrical design of the cell encapsulation pouch eliminates the need for intricate sealing that could lead to sharp edges and rigid connections, which can trigger tissue irritation and fibrosis. The final linear layout of the system simplifies implantation and retrieval through minor incisions using minimally invasive techniques.

[0022] The consistent and programmable oxygen generation from the system was demonstrated in both benchtop and in vivo contexts, enabling adaptability to various cell types and cell dosages. The beneficial impacts of oxygenation on the viability and secretion of insulin were demonstrated in vitro on insulinoma-1 cell (INS-1) aggregates and human pancreatic islets encapsulated at a high density (60,000 islet equivalents (IEQ) / mL). Oxygen generated by the BEAM system was stable, controllable, and sufficient to maintain cell viability and function under hypoxic (1% O₂) conditions in vitro. Additionally, the therapeutic potential of the BEAM system after subcutaneous implantation was evaluated in an allogeneic diabetic rat model, where the oxygenated system implanted subcutaneously reversed diabetes for up to three months without immunosuppression. By comparison, non-oxygenated control animals remained hyperglycemic. Overall, the BEAM system provides a feasible approach to preserving the viability and function of densely packed insulin-secreting cells in the poorly vascularized but clinically attractive subcutaneous site, thus paving the way for development of a practical cell encapsulation system with realistic dimensions and minimally invasive procedures for immunosuppression-free T1D cell replacement therapy. BRIEF DESCRIPTION OF THE DRAWINGS

[0023] FIGs.1A-1C show conceptual design of the BioElectronics-Assisted Macroencapsulation (BEAM) system. The system comprises two integral components: an implantable electrochemical oxygen generator (iEOG) and a core-shell cell encapsulation pouch designed in linear configuration to facilitate minimally invasive implantation, connected by gas- impermeable tubing. FIG.1A is a schematic representation illustrating oxygen generation by electrolytic splitting of tissue moisture. A human sketch is shown to indicate the relative size of a human version system. The power supply needed for electrolysis is not shown. FIG.1B is a representative digital image of a scaled-up BEAM system with a cell encapsulation pouch measuring 15 cm in length and 5 mm in thickness. FIG.1C is a longitudinal section view of the cell encapsulation pouch showing pancreatic islets located in the annular space between an inner silicon tubing and the semipermeable membrane for optimal mass transportation and proximity between cells and the oxygen source. 320349050v3

[0024] FIGs.2A-2G show in situ oxygen generation and biocompatibility of iEOG. FIG.2A is a digital image comparing the dimensions of v1 iEOG and v2 iEOG. FIG.2B shows representative 18-hour measurements of oxygen flow rate (bottom line), current (top line), and voltage (middle line) from a v1 iEOG operated at 10 mA over 2.5 years, collected on day 121, 186, and 879. FIG.2C is a bar graph showing oxygen generation rates of v1 iEOGs operated at 11 mA, measured before and 3 months post-implantation in RNU nude rats (n=4). Data are presented as mean ± SD. Statistical analysis was performed using paired two-tailed Student’s t- test. FIG.2D is a bar graph showing oxygen generation rates of v2 iEOGs operated at 11 mA, measured 2 months post-implantation in RNU nude rats (n=8). Data are presented as mean ± SD. FIG.2E is a bar graph showing oxygen generation rates of v2 iEOGs operated at 11 mA, measured before and 1 month after implantation in immunocompetent SD rats (n=3). Data are presented as mean ± SD. Statistical analysis was performed using paired two-tailed Student’s t- test. FIG.2F shows a representative stereomicroscope image showing a v2 iEOG implanted in the subcutaneous space of SD rats after 1 month. The skin was removed to exposure the implant. FIG.2G shows Masson’s trichrome staining of tissue surrounding the iEOG after 1 month of implantation in SD rats. Arrowheads indicate the presence of blood vessels.

[0025] FIG.3 shows Masson’s trichrome staining demonstrating different levels of fibrotic response in a kinked region (left panel) and intact membrane (right panel) of a Nylon 6 nanofibrous device implanted into intraperitoneal space of C57BL / 6 mice for 1 month. Scale bar: 500 μm.

[0026] FIGs.4A-4H show characterization and scalability of the cell encapsulation pouch. FIG.4A is a schematic illustration for the structure of a cell encapsulation pouch. FIG.4B shows representative digital images showing cell encapsulation pouches in various lengths and diameters, designed to accommodate different cell doses. The inset shows the round end of the pouch. FIG.4C shows SEM images of the pouch surface and cross-section. FIG.4D is a graph showing size distribution of PEBAX fibers. FIG.4E is a graph showing tensile testing of PEBAX cell encapsulation pouches (n=3). FIG.4F shows digital images of a nitinol scaffold and a nitinol-reinforced pouch. Both were bent without kink. FIG.4G shows digital images demonstrating the ability of nitinol-reinforced pouches to reverse to their original structural form following twisting deformation. FIG.4H shows representative microscopy image showing the alginate impregnation onto the encapsulation pouch.

[0027] FIG.5 shows the estimated porosity of electrospun PEBAX membrane. Data are presented as mean ± SD (n=10). The porosity was estimated using ImageJ software based on 10 SEM images taken from 3 independent batches. 320349050v3

[0028] FIG.6 shows Masson’s trichrome staining of heat-sealed nanofibrous nylon 6 devices implanted in the intraperitoneal space of C57BL / 6 mice for 1 month, demonstrating different levels of fibrotic reactions at the heat-sealed end and the middle of the devices (n=4).Scale bar: 1 mm (upper) and 300 μm (lower). Arrowheads ( ) indicate heat-sealed regions.

[0029] FIG.7 shows Masson’s trichrome staining of heat-sealed nanofibrous polyurethanepolyether devices implanted in the intraperitoneal space of C57BL / 6 mice for 1 month, demonstrating different levels of fibrotic reactions at the heat-sealed end and the middle of the devices (n=3). Scale bar: 1 mm (upper) and 400 μm (lower).

[0030] FIG.8 shows Masson’s trichrome staining of heat-sealed nanofibrous PEBAX devices implanted in the intraperitoneal space of C57BL / 6 mice for 1 month, illustrating different levels of fibrotic reactions at the heat-sealed end and the middle of the devices. Scale bar: 100 μm. The membrane was dissolved during the histology process.

[0031] FIG.9 shows the thickness of the fibrotic layers at the end and in center of heatsealed nanofibrous devices made of Nylon 6 (from 4 devices), polyurethane-polyether (from 3 devices), PEBAX (from 4 devices). The data are presented as violin plots. Statistical analyses were performed using unpaired two-tailed Student’s t-test.

[0032] FIG.10 is schematic illustration for preparation of polyethylene glycol template to collect electrospun fibers.

[0033] FIG.11 shows structure of the nitinol braided mesh scaffolds. The scaffold was constructed from 25.4-μm nitinol braided wires, with pore sizes ranging from 100 to 150 μm. These elastic, shape-memory scaffolds effectively maintained the structural integrity of the immuno-protective membrane while minimally impacting mass transport due to their large pore sizes and thin wire thickness.

[0034] FIG.12 shows preparation of INS-1 spheroids using AggrewellTM.

[0035] FIGs.13A-13F show cell loading and encapsulation pouch sealing. FIG.13A is a schematic diagram illustrating stepwise procedure for cell loading and pouch sealing in the BEAM system. FIG.13B shows hematoxylin and eosin (H&E) staining of longitudinal and cross-sections of an encapsulation pouch loaded with INS-1 cell aggregates at 60,000 IEQ / mL. FIG.13C shows bright-field and fluorescence images of the hydrogel layer removed from the encapsulation pouch, containing green fluorescence-labeled INS-1 cell aggregates at 60,000 IEQ / mL. Scale bar: 2 mm. FIG.13D shows a representative digital image of an oxygen transportation tubing infused with a high flow rate of oxygen. FIG.13E shows representative images depicting the connection between the oxygen transportation tubing and the cell encapsulation pouch. The fibrous membrane was removed for visualization purpose. FIG.13F 320349050v3shows the pull-off force measured during the tensile test, indicating the force at which the barb connector is removed from the cell encapsulation pouch (n=5). The data are presented as a min to max box-and-whisker plot.

[0036] FIG.14 shows macroscopic images showing the distribution of 150-μm beads, used as islet mimics, during the injection of suspension into the encapsulation pouch. Gradual dispersion of beads through the pores of the loading tool results in the occupation of interstitial spaces between the loading tool and the encapsulation pouch wall (a transparent tube used for visualization purpose).

[0037] FIGs.15A-15F show the impacts of oxygenation on INS-1 cell aggregates encapsulated in the BEAM system under a hypoxic cell culture condition. FIG.15A is a schematic representation of the experimental setup. FIG.15B is a graph showing operating electrical current and voltage of iEOG recorded during a 24-hour experiment. FIG.15C shows bright-field microscopic images of INS-1 cell aggregates in BEAM systems following a 24-hour incubation period under a hypoxic culture condition (1% oxygen), with and without supplemental oxygenation provided by iEOG. FIG.15D shows dual fluorescence staining of INS-1 cell aggregates within BEAM systems following 24-hour incubation in 1% oxygen, with and without additional oxygen supply from iEOG. FIG.15E shows H&E staining of INS-1 cell aggregates before encapsulation and those loaded in BEAM systems subjected to a hypoxic culture condition (1% oxygen) for 24 hours, with or without oxygen supplementation from iEOG. Grey dashed line represents the interface with the silicone tubing while the black dash line indicates the interface with the fibrous membrane. FIG.15F shows immunofluorescence staining of INS-1 cell aggregates in BEAM systems following 24-hour incubation in 1% oxygen, with and without additional oxygen supply from iEOG.

[0038] FIG.16 shows a longitudinal H&E-stained histological section of a BEAM system encapsulating INS-1 cell aggregates at a density of 60,000 IEQ / mL. The system was subjected to 1% O2 culture condition for 24 hours while the iEOG was operated at 0.27 mA. Scale bars: 400 μm (upper) and 100 μm (lower). The sample was sectioned through the silicone core. No significant difference in the viability were observed between cell aggregates located proximal and distal to the oxygen source.

[0039] FIGs.17A-17F show the impact of oxygenation on viability and function of human pancreatic islets encapsulated in BEAM systems under a hypoxic cell culture condition. FIG. 17A is a schematic representation of the experimental setup. FIG.17B is a graph showing operating electrical current and voltage of iEOG recorded during a 72-hour experiment. FIG. 17C shows dual fluorescence staining of islets encapsulated in BEAM systems subjected to a 24- 320349050v3hour incubation under hypoxic condition (1% oxygen), with and without supplementary oxygenation from iEOGs. Scale bar: 500 µm. Dead cells in bottom right panel are thought to be a cutting artifact from processing the annual hydrogel. FIG.17D shows H&E staining of encapsulated islets after incubation under a normoxic condition and a hypoxic condition, with and without supplementary oxygenation from iEOG. FIG.17E shows immunofluorescence staining of the encapsulated islets after 24-hour incubation under a normoxic and a hypoxic culture condition, with and without supplementary oxygenation from iEOG. FIG.17F is a bar graph showing quantification of insulin secretion from islets encapsulated in BEAM systems, following a 24-hour incubation under normoxic condition (20% oxygen) (n=2) and hypoxic condition (1% oxygen), with and without oxygen supplementation from the iEOG (n=2).

[0040] FIGs.18A-18I show the impact of oxygenation on the metabolic function of pancreatic islets encapsulated within the BEAM system in an allogeneic rat model. FIG.18A is a schematic illustration of experimental design. Current, voltage and battery level were wirelessly spot measured via IR communication. FIG.18B is a digital image demonstrating the BEAM system assembly on a diabetic rat. (1) A pocket on the back of a jacket worn by the rat to secure the iEOG and an electronic box. (2) The oxygen transportation tubing running from iEOG, underneath the jacket to the (3) cell encapsulation pouch implanted in the subcutaneous space of the rat. FIG.18C is a graph showing operating current (top line) and voltage (bottom line) of the iEOG recorded during 88-day study from the electronics memory. FIG.18D is a graph showing cumulative volume of oxygen generated from the iEOGs used on the rats over a 24-hour period (n=4). FIG.18E shows non-fasting blood glucose levels of rats implanted with a cell encapsulation pouch without oxygen supplementation (n=4). FIG.18F shows non-fasting blood glucose levels of rats received implantation of a BEAM system (n=6). FIG.18G shows body weight changes of rats received implants with or without supplemental oxygenation. In color versions of FIGs.18F-18G, black dots represent NBG levels / body weight changes before implantation while green dots represent blood glucose levels / body weight changes during continuous oxygenation, while red dots represent blood glucose levels / body weight changes measured without oxygenation or after the oxygenation cessation. Arrows mark the timepoint of implant retrieval. FIG.18H is a graph showing results of the intraperitoneal glucose tolerance test conducted on healthy rats (n=5), diabetic rats (n=5), rats receiving a BEAM system (n=4; measured on day 35), and rats receiving a cell encapsulation pouch without oxygen supplementary (n=4; measured on day 35). Data are presented as mean ± SD. FIG.18I is a bar graph showing corresponding area under the curve for the groups in FIG.18H. Data are presented as mean ± SD. Statistical analyses were performed using one-way ANOVA. p = 320349050v30.0725 (diabetic rats vs. without oxygenation), p < 0.001 (with oxygenation vs. without oxygenation), p < 0.001 (heathy rats vs. without oxygenation), p = 0.9964 (heathy rats vs. with oxygenation).

[0041] FIG.19 shows workflow in implantation of BEAM device in rats. The electronic controller was programmed to regulate the electrical current, generating an adequate level of oxygen for the islet dose. The electronic controller was then connected to the iEOG on the back of the rats. iEOG performance was wirelessly monitored from outside of the animal cages by an IR receiver linked to a laptop.

[0042] FIG.20 shows a proposed setup for the use of transcutaneous energy transfer system for battery recharge of the BEAM device.

[0043] FIG.21 is a digital image of a rat jacket designed with open flank and abdomen to avoid discomfort.

[0044] FIG.22 is a graph showing percentage body weight change of rats after wearing jackets. The circles indicate the time points that the electronic controllers were placed into the pocket on the back. (n=6).

[0045] FIGs.23A-23B show assessment of transcutaneous exit after implantation of BEAM system. FIG.23A is a digital image showing the tissue healing at the transcutaneous exit. Arrows depict the oxygen transportation tubing. FIG.23B shows Masson’s trichrome staining of the transcutaneous exit, showing tissue integration into the polyester cuffs. No adverse reactions or inflammation were observed surrounding the oxygen transportation tubing. Scale bar: 2 mm.

[0046] FIG.24 shows premature oxygen cessation caused by rats. For implantation design 1, oxygen transportation tubing was damaged by the rats (3 / 3), leading to oxygen leakage. For implantation design 2, rats (2 / 6) escaped from jackets and damaged the electronic components, leading to oxygen cessation for over 24 hours. The first arrows on the graph indicate the time points of oxygen cessation while the second arrows mark the time points of device retrieval.

[0047] FIG.25 shows intraperitoneal glucose tolerance test conducted on healthy rats (n=5), diabetic rats (n=5), rats receiving a BEAM system (n=4; measured on day 35), and rats receiving a cell encapsulation pouch without oxygen supplementary (n=4; measured on day 35). Data are shown for individual subjects.

[0048] FIG.26 shows serum C-peptide analysis of rats receiving devices without oxygenation (n=4) and with oxygenation (n=4) on day 35 post-transplantation before and 90 min after glucose administration. Non-significant differences were found between C-peptide levels of rats received non-oxygenated devices before and 90 min after glucose administration. Higher levels and significant increases in serum C-peptides were observed in rats received oxygenated 320349050v3devices after glucose administration. Statistical analyses were performed using paired two-tailed Student’s t test.

[0049] FIGs.27A-27E show that oxygenation by the BEAM system effectively preserved the viability and function of encapsulated pancreatic islets in an allogeneic rat model. FIG.27A shows stereomicroscopic images showing islet-laden alginate hydrogel layer peeled off from an implant without oxygenation and an implant with oxygenation. FIG.27B shows viability assessment of encapsulated islets by dual fluorescent staining. FIG.27C shows representative immunofluorescence staining of encapsulated islets in implants without oxygen supplementation and implants with oxygen supplementation. FIG.27D shows H&E staining of islets before encapsulation, islets in an implant without oxygenation, and islets in an implant with oxygenation. Similar results were observed among the four rats in non- oxygenation groups and the four rats with continuous oxygenation until retrieval. FIG.27E shows ex vivo GSIS of encapsulated islets in implants without oxygen supplementation (n=3; day 37, 38 and 59) and implants with oxygen supplementation (n=4; day 32, 40, 45, and 88). Data are presented as individual values (left) and mean ± SD (right). Statistical analysis was performed using paired two-tailed Student’s t test.

[0050] FIG.28 shows assessment of islet viability in devices without supplemental oxygenation, with continuous oxygenation, and with oxygen cessation > 24 hours by dual fluorescence staining. Scale bar: 200 μm.

[0051] FIG.29 shows representative H&E staining and immunofluorescence staining of encapsulated islets in implants with oxygenation retrieved on day 32 and day 40 due to the formation of gas phase.

[0052] FIGs.30A-30E show formation of excessive gas surrounding the cell encapsulation pouch. FIG.30A is a digital image showing the location of the gas bubble at the implantation site. FIGs.30B-30D show gas accumulation surrounding devices retrieved from rats with elevated blood glucose levels. The bubbles were released from the tissue upon device retrieval. Black arrows denote gas bubbles, while circled arrows indicate two ends of the cell encapsulation pouch. FIG.30E shows absence of gas phase in devices with oxygen cessation.

[0053] FIG.31 shows a proposed method for scaling up the BEAM system that aims to accommodate 420,000 human IEQ at a loading density of 60,000 IEQ / mL. Seven cell pouches, each measuring 70 mm in length and 7.5 mm in diameter (D) with a capacity of 1.11 mL per pouch, can be arranged in parallel to deliver the required 420,000 human IEQ. The distance (d) between cell pouches is 1 mm. The thickness of cell-laden alginate hydrogel layer (t) is 0.75 mm. The final dimensions of the device, including the iEOG and connectors, are approximately the 320349050v3size of a credit card. A smaller design could be achieved by increasing the loading density, as human islets consume 2-3 times less oxygen compared to rat islets.

[0054] FIGs.32A-32B show a proposed method for scaling up the BEAM devices for delivering small β-cell clusters. As smaller cell clusters (100-150 μm) have been shown to be superior as compared to random-sized pancreatic islets (50-400 μm), it was estimated that device dimensions will house 500,000 β-cell clusters with sizes of 100 μm and 125 μm. The calculations assume that the volume of 1 IEQ is 1.77 nL, while the volume of a 100 μm cell cluster is 0.52 nL and that of a 125 μm cluster is 1.02 nL. This results in a conversion of 60,000 IEQ / mL to 204,230 clusters / mL for 100-μm clusters and 104,117 clusters / mL for 125-μm clusters to achieve an equivalent volume faction. FIG.32A shows a proposed scaled-up device designed to encapsulate 500,000 clusters of 100 μm, which consists of five cell pouches, each 50 mm in length and 5 mm in diameter, with a capacity of 0.5 mL per pouch. These pouches are arranged in parallel. The final dimensions of the device, including the iEOG and connectors, are approximately 29 mm x 68 mm, about 40% of the size of a credit card. FIG.32B shows a proposed scaled-up device to encapsulate 500,000 clusters of 125 μm, which consists of five cell pouches, each measuring 60 mm in length and 7.5 mm in diameter, with a capacity of 0.95 mL per pouch. These pouches are similarly arranged in parallel to accommodate the ~500,000125 μm clusters. The device, including the iEOG and connectors, measures about 41.5 mm x 78 mm, which is roughly 70% of the size of a credit card. For both designs, the distance (d) between cell pouches is 1 mm, and the thickness of the cell-laden alginate hydrogel layer (t) is 0.75 mm.

[0055] FIG.33 shows a proposed design for a fully implantable BEAM system that includes cell pouches encapsulating 500,000 clusters of 100 μm, a rechargeable battery, a transcutaneous energy transfer (TET) charger, and a circuit board. The entire system has a compact footprint of approximately 60 mm × 70 mm—roughly the size of a credit card. Additional batteries can be mounted on the reverse side of the board to extend operational time. A hydrogen diffusion membrane can be integrated into the surface of the electronics housing.

[0056] FIG.34 shows a cell encapsulation pouch of the BEAM device.

[0057] FIG.35 shows heterogeneity of foreign body responses and neovascularization around the devices with and without oxygen supplementation. Scale bar: 500 μm.

[0058] FIG.36 shows experimental setup to measure oxygen and hydrogen flow rates generated from an iEOG implanted in subcutaneous space of rats. 320349050v3DETAILED DESCRIPTION

[0059] The present disclosure relates to implantable cell therapeutic delivery systems, methods of producing these systems, and methods of using the same.

[0060] One aspect of the disclosure relates to an implantable therapeutic delivery system. The implantable therapeutic delivery system includes an electrochemical oxygen generator; a cell compartment comprising a semi-permeable nanofibrous membrane that is impermeable to cellular migration, an internal hydrogel matrix and one or more cells embedded in the hydrogel matrix; and a conduit in fluid communication between the electrochemical oxygen generator and the interior of the cell compartment, wherein the conduit delivers oxygen generated by the electrochemical oxygen generator to the one or more cells embedded in the hydrogel matrix.

[0061] Any of a variety of electrochemical oxygen generators can be used in the disclosed implantable therapeutic delivery system. One example is described in U.S. Patent No.6,368,592 to Colton et al., which is hereby incorporated by reference in its entirety. This oxygen generator electrolyzes water to provide oxygen that can be supplied to cells in vitro or in vivo, but without generating free hydrogen due to the presence of a multilayer electrolyzer sheet having a proton exchange membrane sandwiched by an anode layer and a cathode layer. Another example is described in U.S. Patent No.10,231,817 to Tempelman et al., which is hereby incorporated by reference in its entirety. This oxygen generator can either be an externally worn oxygen concentrator or an implanted electrolysis devices that utilizes water from body fluids to provide oxygen that can be supplied to cells in vivo. A further example includes an implantable water electrolyzer as described in U.S. Patent No.11,773,496 to Schwenk et al., which is hereby incorporated by reference in its entirety. This electrolytic gas generator may comprise a membrane electrode assembly (MEA), an anode support, a cathode support, an anode current collector assembly, and a cathode current collector assembly to produce oxygen gas.

[0062] In an exemplary device illustrated in FIGs.1A-1C, an iEOG of the type manufactured by Giner Inc. is illustrated. In these iEOGs, the electrolyzer components are housed within a medical-grade titanium enclosure with a porous polymeric membrane integrated onto one face, functioning as a window to harvest interstitial water vapor from surrounding tissue for electrolysis. The iEOGs can be controlled to a settable current with corresponding voltages from 1.4 to 1.8 V, in a range suitable for the splitting of water into hydrogen and oxygen, which were released through two separate outlets (FIG.1A). The oxygen outlet delivers gas by convection to an oxygen-permeable tubing traversing the cylindrical cell encapsulation pouch, enabling oxygen diffusion along its length to the encapsulated cells (FIGs.1B-1C). An alginate-impregnated fibrous membrane can be used as the immuno-protective barrier, which allows for selective 320349050v3diffusion of nutrients and any secreted therapeutic agents while preventing the infiltration of host cells.

[0063] In certain embodiments, the conduit comprises a first portion that is impermeable to oxygen and a second portion that is permeable to oxygen.

[0064] In certain embodiments, the first portion is primarily located externally of the cell compartment and the second portion is entirely located internally to the cell compartment. This ensures that oxygen generated by the electrochemical oxygen generator is delivered to the cell compartment. Exemplary materials that can be used for the first portion (oxygen impermeable) of the conduit include, without limitation, polytetrafluoroethylene, TYGON®(Saint-Gobain Corporation), polypropylene, and polycarbonate. Any other oxygen impermeable materials that can be appropriately coupled to the second portion of the conduit can also be used.

[0065] In certain embodiments, the second portion of the conduit is located centrally within the cell compartment and extends the full length of the cell compartment. Exemplary gas permeable materials include, without limitation, silicone rubber and expanded PTFE having a pore size of 0.5 μm or less. Any other gas permeable materials that can be appropriately coupled to the first portion of the conduit can also be used.

[0066] In certain embodiments, a terminal end of the second portion of the conduit is sealed such that oxygenation of cells in the device occurs as a result of oxygen diffusing through the second portion into the hydrogel matrix in which the one or more cells are embedded.

[0067] In certain embodiments, the implantable therapeutic delivery system further comprises a hermetical seal formed at the junction of the cell compartment membrane, and the first and second portions of the conduit. As demonstrated in the accompanying examples, a pair of mating connectors were employed in combination with an adhesive. The pair of mating connectors included (i) a polyurethane ring that was physically coupled to the cell encapsulating pouch during its construction via electrospinning, and (ii) a polycarbonate barb connector that was incorporated onto the oxygen-impermeable tubing. Alternative materials can, of course, be substituted for the polyurethane and polycarbonate. Regardless of the materials used, the cell encapsulation pouch can be sealed by ensuring that the barb connector fits securely within the polyurethane ring of the cell encapsulation pouch. This mechanical connection can optionally be reinforced using a suitable adhesive such as cyanoacrylate adhesive.

[0068] In certain embodiments, the semi-permeable nanofibrous membrane is formed by electrospinning. Suitable methods for making the semi-permeable nanofibrous membrane are described in WO / 2025 / 106977 to Ma et al., which is hereby incorporated by reference in its entirety. 320349050v3

[0069] In certain embodiments, the semi-permeable nanofibrous membrane is formed of nanofibers comprising one or more medical-grade elastomers. Suitable medical-grade elastomers that can be used include, without limitation, the ones selected from the group of polyethylene glycol, polyamide, polyether block amide, polycarbonate urethane, thermoplastic silicon- polycarbonate-urethane, polyether urethane, and combinations thereof.

[0070] In certain embodiments, the semi-permeable nanofibrous membrane is seamless. Seamless construction of the nanofibrous membrane is described in the accompanying examples.

[0071] In certain embodiments, the semi-permeable nanofibrous membrane further comprises one or more nitinol mesh layers.

[0072] In certain embodiments, the semi-permeable nanofibrous membrane is either (i) a hydrogel-reinforced nanofibrous membrane, (ii) a nitinol mesh-reinforced nanofibrous membrane, or (iii) a hydrogel and nitinol mesh-reinforced nanofibrous membrane. Suitable semi- permeable nanofibrous membranes that can be used include, but are not limited to, the ones described in WO / 2025 / 106977 to Ma et al., which is hereby incorporated by reference in its entirety.

[0073] In certain embodiments, the hydrogel material can comprise a natural polymeric material, a synthetic polymeric material, or a combination thereof.

[0074] Suitable natural polymeric materials that can be used include, without limitation, collagen, hyaluronate, fibrin, alginate, agarose, chitosan, bacterial cellulose, elastin, keratin, derivatives thereof, and combinations thereof. In some embodiments, the hydrogel material comprises a pure alginate, a modified alginate, or a mixture of pure and modified alginate. In some embodiments, the modified alginate is a zwitterionically modified alginate.

[0075] Suitable synthetic polymeric materials that can be used include, without limitation, polyethylene glycol (PEG), poly(acrylic acid), poly(ethylene oxide), poly(vinyl alcohol), polyphosphazene, poly(hydroxyethyl methacrylate), triazole-zwitterion hydrogels (TR-qCB, TR- CB, TR-SB), poly(sulfobetaine methacrylate), carboxybetaine methacrylate, poly[2- methacryloyloxyethyl phosphorylcholine, N-hydroxyethyl acrylamide, a copolymer thereof, a derivatives thereof, and a combination thereof.

[0076] In certain embodiments, the nitinol mesh-reinforced nanofibrous material comprises one or more nanofiber layers, such as first and second nanofibrous layers with the nitinol mesh sandwiched between the first and second nanofibrous layers, optionally wherein fibers of the first and second nanofibrous layers are bonded to one another via pores in the nitinol mesh.

[0077] In some embodiments, the nitinol mesh-reinforced hydrogel material has a thickness of about 10 to about 200 μm, about 10 to about 100 μm, about 10 to about 50 μm, about 50 to 320349050v3about 100 μm, about 100 to about 150 μm, or about 150 to about 200 μm, such as about 10 to about 20 μm, about 20 to about 30 μm, about 30 to about 40 μm, about 40 to about 50 μm, about 50 to about 60 μm, about 60 to about 70 μm, about 70 to about 80 μm, about 80 to about 90 μm, about 90 to about 100 μm, about 100 to about 110 μm, about 110 to about 120 μm, about 120 to about 130 μm, about 130 to about 140 μm, about 140 to about 150 μm, about 150 to about 160 μm, about 160 to about 170 μm, about 170 to about 180 μm, about 180 to about 190 μm, or about 190 to about 200 μm.

[0078] In certain embodiments, the hydrogel matrix is a natural polymeric material, a synthetic polymeric material, or a combination thereof.

[0079] In certain embodiments, the cells positioned within the hydrogel matrix secrete at least one therapeutic agent.

[0080] In certain embodiments, the implantable therapeutic delivery system further comprises a second therapeutic agent positioned within the hydrogel material.

[0081] In certain embodiments, the preparation of cells comprises a preparation of single cells or a preparation of cell aggregates.

[0082] In certain embodiments, the preparation of cells comprises a preparation of primary cells or a preparation of immortalized cells.

[0083] In some embodiments, the preparation of cells can include mammalian cells. Suitable mammalian cells that can be used include, without limitation, primate cells, rodent cells, canine cells, feline cells, equine cells, bovine cells, and porcine cells. In some embodiments, the preparation of cells includes human cells.

[0084] In some embodiments, the preparation of cells can include stem cells or stem cell derived cells. The stem cells can be pluripotent, multipotent, oligopotent, or unipotent stem cells. Suitable stem cells that can be used include, without limitation, embryonic stem cells, epiblast cells, primitive ectoderm cells, primordial germ cells, and induced pluripotent stem cells. Alternatively, the preparation of cells can include cells that are derived from such stem cells, examples of which are described below.

[0085] In some embodiments, the preparation of cells can include, one or more of smooth muscle cells, cardiac myocytes, platelets, epithelial cells, endothelial cells, urothelial cells, fibroblasts, embryonic fibroblasts, myoblasts, chondrocytes, chondroblasts, osteoblasts, osteoclasts, keratinocytes, hepatocytes, bile duct cells, islet cells, thyroid, parathyroid, adrenal, hypothalamic, pituitary, ovarian, testicular, salivary gland cells, adipocytes, embryonic stem cells, mesenchymal stem cells, neural cells, endothelial progenitor cells, hematopoietic cells, precursor cells, mesenchymal stromal cells, Baby Hamster Kidney (BHK) cells, Chinese 320349050v3Hamster Ovary cells, Human Amniotic Epithelial (HAE) cells, choroid plexus cells, chromaffin cells, adrenal chromaffin cells, pheochomocytoma cell line PC12, human retinal pigment epithelium cells, recombinant human retinal pigment epithelium cells, NGF-secreting Baby Hamster Kidney (BHK) cells, human bone marrow-derived stem cells transfected with GLP-1, BDNF-producing fibroblasts, NGF-producing cells, CNTF-producing cells, BDNF-secreting Schwann cells, IL-2-secreting myoblasts, endostatin-secreting cells, and cytochrome P450 enzyme overexpressed feline kidney epithelial cells, myogenic cells, embryonic stem cell- derived neural progenitor cells, irradiated tumor cells, proximal tubule cells, neural precursor cells, astrocytes, genetically engineered cells.

[0086] In some embodiments, the preparation of cells includes a preparation of islet cells that release insulin and glucagon (as the therapeutic agents). As used herein, the term islet cells refers generically to one or more individual cell types that are present in islets as well as islets, which contain a population of different cell types including alpha, beta, delta, and gamma hormone secreting cells as well as cells associated with blood vessels and ducts.

[0087] In some embodiments, the preparation of islet cells can include a preparation of alpha cells, a preparation of beta cells, a preparation of delta cells, a preparation of gamma cells, or combinations thereof, which are not present in the form of a primary isolated islet per se. The preparation of islet cells may originate from islets, i.e., be isolated therefrom, or the preparation of islet cells may be derived from stem cells that are cultured under conditions suitable to cause differention into alpha cells, beta cells, delta cells, and / or gamma cells. Combinations of alpha cells, beta cells, delta cells, and / or gamma cells can be aggregated into islet-like clusters or aggregates.

[0088] In some embodiments, the preparation of islet cells can be a preparation of islets, e.g., human, porcine, or rodent islets. The preparation of islets is harvested from pancreatic tissue and cultured under appropriate conditions prior to use in accordance with the present invention. The preparation of islets may comprise a density between about 1x103to 2x106islet equivalents (IEQs) / mL. For example, the preparation of islets can comprise a density ranging between about 1x103up to about 5x103, 1x103up to about 1x104, 1x103up to about 5x104, 1x103up to about 1x105, 1x103up to about 5x105, 1x103up to about 1x106, 5x103up to about 1x104, 5x103up to about 5x104, 5x103up to about 1x105, 5x103up to about 5x105, 5x103up to about 1x106, 5x103up to about 2x106, 1x104up to about 5x104, 1x104up to about 1x105, 1x104up to about 5x105, 1x104up to about 1x106, 1x104up to about 2x106, 5x104up to about 1x105, 5x104up to about 5x105, 5x104up to about 1x106, 5x104up to about 2x106, 1x105up to about 5x105, 1x105up to about 1x106, 1x105up to about 2x106, 5x105up to about 1x106, 5x105up to about 2x106, or 1x106320349050v3up to about 2x106islet equivalents (IEQs) / mL. The above densities are based on the density of the liquid that is loaded into the cell compartment.

[0089] The therapeutic agent and / or the second therapeutic agent can include one or more cell factors or biologically active agents to enhance cell growth, differentiation, and / or survival of the cells positioned within the hydrogel matrix. Suitable biologically active agents include, without limitation, a protein, peptide, antibody or antibody fragment thereof, antibody mimetic, a nucleic acid, a small molecule, a hormone, a growth factor, an angiogenic factor, a cytokine, an anti-inflammatory agent, an anti-fibrotic agent, and combinations thereof.

[0090] Exemplary growth factors include, without limitation, fibroblast growth factors (FGFs) such as FGF1, FGF4, FGF19, and FGF21; nerve growth factors (NGFs), epiderma growth factors (EGFs), transforming growth factors, hepatocyte growth factors (HGFs), platelet- derived growth factors (PDGFs), insulin-like growth factors (IGFs), IGF binding proteins, basic fibroblast growth factors, and vascular endothelial growth factors (VEGF).

[0091] Exemplary angiogenic factors include, without limitation, VEGF, bFGF, HGF, PDGF, ANG-1, and IGF-1.

[0092] Exemplary cytokines include, without limitation, interleukins, lymphokines, monokines, colony stimulating factors, chemokines, interferons and tumor necrosis factor (TNF).

[0093] Exemplary anti-inflammatory agents include, without limitation corticosteroids such as prednisone, cortisone, and methylprednisolone; and non-steroidal anti-inflammatory agents (NSAIDs) such as ibuprofen, naproxen, celecoxib, diclofenac, indomethacin, oxaprozin, and piroxicam.

[0094] Exemplary antifibrotic agents include, without limitation, nintedanib, GW2580 (the cFMS Receptor Tyrosine Kinase Inhibitor 5-[[3-methoxy-4-[(4-methoxyphenyl)methoxy]- phenyl]methyl]pyrimidine-2,4-diamine), and pirfenidone. In some embodiments, the anti-fibrotic agent can be in crystalline form. While several of these agents have previously been evaluated for preventing fibrotic responses against intraperitoneally inserted alginate microcapsules (see Farah et al., "Long-term Implant Fibrosis Prevention in Rodents and Non-human Primates Using Crystallized Drug Formulations," Nat. Mater., 18(8):892-904 (2019); PCT Publ. No. WO 2017 / 176804, each of which is hereby incorporated by reference in its entirety), these prior reports raise several potential problems. The inability to reliably retrieve all capsules raises safety concerns and issues associated with the regulatory approval process. Because the microcapsules are randomly distributed in the intraperitoneal cavity, the drug concentration is highly varied and depends on the density of capsules. Thus, it is difficult to reliably control the drug concentration in the vicinity of all capsules within the therapeutic window. Finally, co- 320349050v3encapsulation of cells and crystalline drugs in microcapsules raises concern that the empty space left after the drug dissolves may devastate the integrity of any immuno-isolation barrier. Indeed, others have used degradable materials to release pirfenidone for the treatment of corneal abrasion (Tawfik et al., "Dual Drug-loaded Coaxial Nanofibers for the Treatment of Corneal Abrasion," Internat’l J Pharmaceutics, 581:119296 (2020), which is hereby incorporated by reference in its entirety), but the degradable materials precluded immuno-isolation, afforded limited drug- loading capacity, and achieved a fairly short (< 10 h) drug release period that was incompatible with long-term delivery.

[0095] The cell encapsulating pouch can be prepared by an electrospinning technique using medical grade polymers as a base material (FIG.4A). By way of example, medical-grade polyether block amide (Arkema Pebax™) can be used to prepare the cell encapsulating pouch. The pouch can be sealed using conventional sealing methods, such as heat, ultrasonic welding, or adhesives.

[0096] To minimize the area requiring sealing, an edge-free cell pouch can be used, such as an edge-free round-ended pouch (FIG.10). To fabricate round-ended, edge-free cell encapsulation pouch, a PEG template of the desired shape can be used. A medical-grade polyurethane ring can be affixed to one end of the PEG template, functioned as a part of the barb connector to seal the pouch. Once the pouch is prepared, the PEG template can be dissolved using water.

[0097] To impart the immuno-protective property, the cell encapsulating pouch can be impregnated by soaking in a solution that will facilitate coating of the cell encapsulating pouch with a cross-linking agent. By way of example, a barium chloride solution can be used to promote cross-linking of an alginate solution. Once the cell encapsulating pouch is coated with cross-linking agent, cells can be loaded through a loading apparatus, into the pouch. Then the loading apparatus can be withdrawn, and the oxygen-transportation tubing can be inserted. The sealed cell encapsulation pouch can be then connected to an iEOG.

[0098] Alternatively, the cell encapsulating pouch can be first soaked in a solution impart the immuno-protective property without the crosslinking agent. By way of example, a solution of ultrapure alginate (SLG100) can be used. Once the cell encapsulating pouch is coated with a layer of immuno-protective coating, cells can be loaded through the loading apparatus, into the pouch. Then the cell encapsulating pouch can be immersed in a crosslinking solution. By way of example, a solution of calcium chloride and barium chloride can be used that will cause the divalent ions to diffuse through the porous membrane of the cell encapsulating pouch, initiating 320349050v3alginate gelation. Then the loading apparatus can be withdrawn, and the oxygen-transportation tubing can be inserted. The sealed cell encapsulation pouch can be then connected to an iEOG.

[0099] Once the implantable therapeutic delivery system has been completely assembled, the delivery system can then be implanted in a subject for delivery of therapeutic agent(s) to the subject while the device remains implanted.

[0100] In particular, the devices can be implanted using laproscopic surgical procedures or open surgical sites, and the devices can be placed subcutaneously, transcutaneously, preperitoneally, transperitoneally, or intraperitoneally. In some embodiments, implanting involves suturing the device or system to a body wall of the subject; anchoring the device to a body wall of the subject via a transabdominal portal; wrapping the delivery device or system in omentum of the subject; positioning the device in a cavity between the liver and the diaphragm; or anchoring the device to the diaphragm. Unanchored implantation is also contemplated.

[0101] Both veterinary and medical uses are contemplated. Thus, exemplary subjects include, without limitation, a human, a mouse, a rat, a dog, a cat, a pig, a sheep, a cow, a horse, and a nonhuman primate.

[0102] By implanting the implantable therapeutic delivery system, the systems can be used to deliver a therapeutic agent to a subject in need thereof. The treatment by implantation can be carried out for a limited duration over a period of days, weeks, or months. Thus, it is also contemplated that the method of treatment further involves retrieving the implantable cell containing device from the subject when no longer needed or when the device needs replacement, and optionally implanting a replacement implantable therapeutic deliver system after the initial device is retrieved. It is also contemplated that the method of treatment further involves replacing a battery of the electrochemical oxygen generator without retrieving the implantable therapeutic delivery system.

[0103] The introduction of one or more contrast agents allows for monitoring of the device. Methods of in vivo monitoring include but are not limited to confocal microscopy, 2-photon microscopy, high frequency ultrasound, optical coherence tomography (OCT), photoacoustic tomography (PAT), computed tomography (CT), magnetic resonance imaging (MRI), single photon emission computed tomography (SPECT), and positron emission tomography (PET). These alone or combined can provide useful means to monitoring the implantable device. Monitoring of the device may be used to determine when to remove and replace a device, as necessary.

[0104] According to one embodiment, the subject has diabetes, is in need of diabetes treatment, and the method of treating diabetes in the subject involves implanting an implantable 320349050v3therapeutic delivery system as described herein. In some embodiments, the hydrogel matrix comprises a preparation of cells that release insulin, glucagon, or a combination thereof. Exemplary preparation of cells include one or more of islet cells, primary islets isolated from pancreas, and islet cells or islet-like aggregates derived from a preparation of stem cells such as pluripotent, multipotent, oligopotent, or unipotent stem cells, including embryonic stem cells, epiblast cells, primitive ectoderm cells, primordial germ cells, and induced pluripotent stem cells. Exemplary preparation of islets is a preparation of primate islets, rodent islets, canine islets, feline islets, equine islets, bovine islets, or porcine islets.

[0105] According to one embodiment, the subject has a bleeding disorder, is in need of treatment for the bleeding disorder, and the method of treating a bleeding disorder in a subject involves implanting an implantable therapeutic delivery system as described herein. In accordance with this embodiment, the bleeding disorder can be any bleeding disorder, such as hemophilia A, hemophilia B, von Willebrand disease, Factor I deficiency, Factor II deficiency, Factor V deficiency, Factor VII deficiency, Factor X deficiency, Factor XI deficiency, Factor XII deficiency, and Factor XIII deficiency. In some embodiments, the hydrogel matrix comprises a preparation of cells that release one or more blood clotting factors selected from the group of Factor I, Factor II, Factor V, Factor VII, Factor VIII, Factor IX, Factor X, Factor XI, Factor XII, Factor XIII, and combinations thereof. Exemplary preparation of cells include recombinant myoblasts, mesenchymal stromal cells, endothelial cells, induced pluripotent stem cell derived endothelial cells, induced pluripotent stem cell derived mesenchymal stromal cells, and a combination thereof.

[0106] In another embodiment, the subject has a lysosomal storage disorder, is in need of treatment for the lysosomal storage disorder, and the method treating a lysosomal storage disease in a subject involves implanting an implantable therapeutic delivery system as described herein into the subject having the lysosomal storage disorder. In accordance with this embodiment, the hydrogel matrix comprises a preparation of cells that release an enzyme selected from the group of α-L-iduronidase, Iduronate-2-sulfatase, α-glucuronidase, Arylsulfatase A, alpha-Galactosidase A, and combinations thereof. Exemplary preparation of cells include one or more of hematopoietic stem cells, fibroblasts, myoblasts, Baby Hamster Kidney (BHK) cells, Chinese Hamster Ovary cells, Human Amniotic Epithelial (HAE) cells, mesenchymal stromal cells, induced pluripotent stem cell derived mesenchymal stromal cells, and combinations thereof.

[0107] According to one embodiment, the subject has a neurological disorder, is in need of treatment for the neurological disorder, and the method of treating a neurological disorder in a subject involves implanting an implantable therapeutic delivery system as described herein. In 320349050v3accordance with this embodiment, the neurological disorder is Parkinson’s disease, Alzheimer’s disease, epilepsy, Huntington’s disease, Amyotrophic lateral sclerosis, chronic pain, a sensory disorder such as visual loss, hearing loss, peripheral nerve injury, and spinal cord injury, and the hydrogel matrix comprises a preparation of cells that release a molecule selected from the group of cerebrospinal fluid, extracellular fluid, levodopa, nerve growth factor (NGF), ciliary neurotrophic factor (CNTF), BLP-1, brain-derived neurotrophic factor (BDNF), vascular endothelial growth factor (VEGF), enkephalin, adrenaline, catecholamine, and combinations thereof. The preparation of cells comprises choroid plexus cells, chromaffin cells, pheochomocytoma cell line PC12, human retinal pigment epithelial cells, NGF-secreting Baby Hamster Kidney (BHK) cells, myoblasts, human bone marrow-derived stem cells transfected with GLP-1, BDNF-producing fibroblasts, NGF-producing cells, CNTF-producing cells, adrenal chromaffin cells, BDNF-secreting Schwann cells, myogenic cells, embryonic stem cell-derived neural progenitor cells, and combinations thereof.

[0108] In another embodiment, the subject has a cancerous condition, is in need of treatment for the cancerous condition, and the method of treating a cancer in a subject involves implanting an implantable therapeutic deliver system as described herein into the subject having the cancerous condition. In accordance with this embodiment, the hydrogel matrix comprises a preparation of cells that release a molecule selected from IL-2, endostatin, cytochrome P450 enzyme, a tumor antigen, a cytokine, and combinations thereof. The preparation of cells comprises IL-2-secreting myoblasts, endostatin-secreting cells, Chinese Hamster Ovary cells, cytochrome P450 enzyme overexpressed feline kidney epithelial cells, irradiated tumor cells, and combinations thereof.

[0109] In another embodiment, the subject has a chronic eye disease, is in need of treatment for the chronic eye disease, and the method of treating a chronic eye disease in a subject involves implanting an implantable therapeutic delivery system as described herein into the subject having the chronic eye disease. The chronic eye disease may be any one of age-related macular degeneration, diabetic retinopathy, retinitis pigmentosa, glaucoma, macular telangiectasia, and combinations thereof. In accordance with this embodiment, the hydrogel matrix comprises a preparation of cells that release a molecule selected from ciliary neurotrophic factor, antagonists against vascular endothelial growth factor and platelet-derived growth factor, and combinations thereof. The preparation of cells comprises human retinal pigment epithelium cells, recombinant human retinal pigment epithelium cells, and combinations thereof.

[0110] In one embodiment, the subject has kidney disease (kidney failure), is in need of treatment for the kidney disease (kidney failure), and the method of treating a kidney failure in a 320349050v3subject involves implanting an implantable therapeutic delivery system as described herein into the subject having the kidney disease (kidney failure). In accordance with this embodiment, the hydrogel matrix comprises a preparation of cells that release a therapeutic molecule suitable for treating the kidney failure. The therapeutic molecule can be dopamine, atrial natriuretic peptide, and combinations thereof. The hydrogel matrix comprises a preparation of renal proximal tubule cells, mesenchymal stem cells, and combinations thereof.

[0111] In one embodiment, the subject has chronic pain, is in need of treatment for the chronic pain, and the method of treating a chronic pain in a subject involves implanting an implantable therapeutic delivery system as described herein into the subject having the chronic pain. The chronic pain can be any chronic pain condition including, without limitation, those caused by a degenerative joint (e.g., back, hip, or knee), peripheral neuropathy, or cancer. In accordance with this embodiment, the hydrogel matrix comprises a preparation of cells that release a molecule selected from the group consisting of catecholamine, opioid peptides, enkephalins, and combinations thereof. The preparation of cell comprises chromaffin cells, neural precursor cells, mesenchymal stem cells, astrocytes, and genetically engineered cells, and combinations thereof.

[0112] Another aspect of the disclosure relates to a method of forming a seamless nanofibrous membrane. This method involves forming an enlarged diameter support about a mandrel, the support being formed of a dissolvable material; and electrospinning a nanofibrous membrane about the enlarged diameter support and a portion of the mandrel.

[0113] In some embodiments, the method further involves dissolving the enlarged diameter support; and removing the seamless nanofibrous membrane from the mandrel.

[0114] Another aspect of the disclosure relates to a method of making a cell encapsulation system suitable for implant. This method involves providing a semi-permeable nanofibrous membrane structure; inserting a permeable conduit into the semi-permeable nanofibrous membrane, the permeable conduit having an unsealed end; and introducing cells and hydrogel precursor via the permeable conduit, whereby the cells and hydrogel precursor are forced into the space between the permeable conduit and the nanofibrous membrane, followed by crosslinking of the hydrogel precursor to form the hydrogel matrix. This method further involves joining together the unsealed end of the permeable conduit to a distal end of an impermeable conduit coupled to an electrochemical oxygen generator; and sealing the semi-permeable nanofibrous membrane about the distal end of the impermeable conduit.

[0115] In some embodiments, the semi-permeable nanofibrous membrane structure has a single opening, and the permeable conduit is inserted through the single opening. 320349050v3

[0116] In some embodiments, sealing is carried out by adhesively sealing the semi- permeable nanofibrous membrane about the distal end of the impermeable conduit. In other embodiments, sealing is carried out by installing a hermetic coupling between the semi- permeable nanofibrous membrane and the distal end of the impermeable conduit.

[0117] Another aspect of the disclosure relates to a device for forming an implantable therapeutic delivery system. This device comprises an electrochemical oxygen generator; a compartment comprising a semi-permeable nanofibrous membrane that is impermeable to cellular migration; a conduit in fluid communication between the electrochemical oxygen generator and the interior of the compartment, wherein the conduit delivers oxygen generated by the electrochemical oxygen generator to the interior of the compartment.

[0118] Another aspect of the disclosure relates to a kit. This kit comprises the device as described herein; one or more precursors for forming a hydrogel matrix; and instructions for loading a composition comprising cells and the one or more precursors into the compartment of the device, and for forming the hydrogel matrix in situ.

[0119] Wherever the word “about” is employed herein in the context of dimensions (e.g. distances, sizes), time, amounts (relative amounts, concentration, etc.), cell densities, etc., it will be appreciated that such variables are approximate and as such may vary by ± 10%, for example ± 5% and preferably ± 2% (e.g., ± 1%) from the numbers specified herein. EXAMPLES

[0120] The examples below are intended to exemplify the practice of embodiments of the disclosure but are by no means intended to limit the scope thereof. Materials and Methods

[0121] Preparation of Rat INS-1 Cell Aggregates: INS-1832 / 13 cell aggregates were prepared using a commercially available microwell plate (Aggrewell; STEMCELL Technologies). Briefly, 5.9 x 106INS-1832 / 13 cells were added into a well of an AggreWellTM400 plate, followed by centrifugation at 100 x g for 3 minutes. After 48-hour incubation, the cell aggregates were harvested and cultured in Roswell Park Memorial Institute medium (RPMI- 1064 GlutaMAXTM; Gibco) supplemented with fetal bovine serum (FBS; 10% (v / v); Hyclone), HEPES (10 mM; Gibco), sodium pyruvate (1 mM; Gibco), penicillin (100 IU / mL; Gibco), streptomycin (100 µg / mL; Gibco), and β-mercaptoethanol (50 µM; Gibco).

[0122] Human Pancreatic Islet: Human islets (Purity 80-95%) were provided by Dr. C. Liu from the Human Islet Core within the University of Pennsylvania, where the islets were isolated 320349050v3according to the guidelines outlined by the Clinical Islet Transplantation (CIT) consortium protocols. These islets were then transported in cold CIT culture media and cultured for 1-3 days at 37oC before encapsulation.

[0123] Rat Pancreatic Islet Isolation: Rat pancreatic islets were isolated from Lewis rats (200-250 grams; Charles River Laboratories) using a previously established enzyme digestion procedure (Pham et al., “Surface-Triggered in Situ Gelation for Tunable Conformal Hydrogel Coating of Therapeutic Cells and Biomedical Devices,” Advanced Functional Materials 31(21):2010169 (2021); Nguyen et al., “Engineering “Cell-Particle Hybrids” of Pancreatic Islets and Bioadhesive FK506-Loaded Polymeric Microspheres for Local Immunomodulation in Xenogeneic Islet Transplantation,” Biomaterials 221:119415 (2019); Tran et al., “Prolongation of Graft Survival via Layer-By-Layer Assembly of Collagen and Immunosuppressive Particles on Pancreatic Islets,” Biomaterials 290:121804 (2022), which are hereby incorporated by reference in their entirety). First, 10 mL of LiberaseTMTL (0.15%; Research grade, Roche) in ice-cold Hank’s balanced salt solution (HBSS; Corning) was slowly infused into the pancreas through the pancreatic duct. The pancreas was then incised from associated body parts and incubated at 37oC for 31 minutes. Then, 50 mL of ice-cold RPMI-1640 supplemented with 10% FBS was added to terminate the enzymatic digestion. The resulting tissue pellet was washed with RPMI-1640 and filtered through a 450 µm sieve. Islets were purified by a density gradient centrifugation using a Histopaque 1077 / RPMI-1640 media gradient. Finally, the islets were manually picked under a light microscope (SZ61, Olympus) to achieve higher purity. Preparation and Assembly of the BEAM System

[0124] Manufacture and Sterilization of iEOGs: The iEOGs were designed and GMP- manufactured by Giner Inc. Sterilization of the iEOGs was performed using the nitrogen dioxide method (Noxilizer Inc). Before integration with a cell encapsulation pouch, the operational efficacy of the sterilized iEOGs was evaluated using a highly sensitive mass flow meter (AlicatTMScientific MW-0.5SSCCM-D). Only iEOGs with oxygen production levels within 85% to 115% of their theoretical capability were deemed suitable for use in the experimental procedures.

[0125] Regulation of Oxygen Generation: The rate of oxygen generation (Q) is regulated by the modulation of the electrical current (I) applied to the system, as the oxygen flow rate is directly proportional to the current, according to Faraday's law of electrolysis with adjustments for factors such as temperature, back pressure, and internal losses within the electrolyzer. The equation governing oxygen production is as follows: 320349050v3where Vmis the molar volume of oxygen under standard ambient temperature and pressure (SATP: 25°C and 1 atm); F is Faraday’s constant derived from the product of the elementary charge (e) and Avogadro’s number (NA); and z indicates the number of electrons required to produce one molecule of oxygen, typically 4 electrons per molecule of oxygen.

[0126] A custom-built electronic controller was developed by Giner Inc to regulate and maintain a constant current circuit. When the electrolyzer current was set from 0.2 mA to 12 mA, the corresponding voltage required to maintain the set point current was from 1.4 V to 1.8 V, which falls within the expected range for electrolysis.

[0127] In Vitro Assessment of iEOG Performance: The long-term in vitro performance of implantable electrochemical oxygen generators (iEOGs) was evaluated in normal saline. Three iEOGs were configured to operate at a current of 10 mA using a custom electronic controller developed by Giner Inc for 11 months, 2 years, and 2.5 years. Current and corresponding voltage were measured every 30 s throughout the study. On a weekly basis, each unit was connected for an 18-hour period to a sensitive mass flow meter, calibrated at the factory for oxygen measurement (Whisperlite, Model MW-0.5SSCCM-D, Alicat) to measure the oxygen flow rate and demonstrate it corresponded as expected to the current setpoint.

[0128] In Vivo Assessment of iEOG Performance: To assess the in vivo oxygenation capability, sterile iEOGs were implanted into the subcutaneous space of immunodeficient RNU nude rats or immunocompetent SD rats and operated at 11 mA. The flow rates of oxygen generated from these iEOGs were recorded prior to implantation and on the day of explantation. For the experiment on RNU nude rats, the oxygen generation rates were measured from the iEOGs in tissue samples collected post-mortem. For the experiment on SD rats, a flow meter (Whisperlite, Model MW-0.5SSCCM-D, Alicat) was directly connected to the oxygen outlet of the iEOG exposed through the skin. See FIG.36.

[0129] Preparation of Cell Encapsulation Pouch: The fibrous cell encapsulation pouch was prepared using an electrospinning method. Specifically, medical-grade polyether block amide (Arkema Pebax™) was dissolved in 1,1,1,3,3,3-hexafluoro-2-propanol (HFIP) at a concentration of 12.5% (w / v) and electrospun at 14 kV through a 23-gauge blunt needle, which served as the spinneret. The polymer solution was fed at a flow rate of 1.5 mL / h, with the spinneret-to- mandrel distance set to 11 cm. To fabricate round-ended, edge-free cell encapsulation pouch, a PEG template of the desired shape was deposited onto a conductive mandrel by injection 320349050v3molding to collect electrospun fibers. For the rat scale pouch, a two-part 3D-printed mold was fabricated using a Formlabs 3 printer (Formlabs Inc) with Clear V4 photopolymer resin. The mold was assembled onto one end of a steel rod with a diameter of 1.6 mm, and melted PEG was introduced into central cavity within the mold. After allowing the PEG to solidify at room temperature for 10 minutes, the mold was disassembled, resulting in a 700-µm-thick PEG layer attached to one end of the rod. A 1-cm medical-grade polyurethane ring (Scientific Commodities Inc) was affixed to one end of the PEG template, functioned as a part of the barb connector to seal the pouch. For long pouches, a braided nitinol mesh scaffold (medical grade; Secant Group) with a diameter matching that of the PEG template was first immersed in a 1.5% (w / v) PEBAX solution in HFIP and subsequently positioned over the PEG template. The mandrel was then mounted onto a rotator operating at 450 rpm to facilitate the collection of electrospun fibers for 10 minutes. Afterward, the mandrel was removed and immersed in distilled water for 30 minutes to completely dissolve the PEG template. The resulting pouch was washed 5 times with distilled water and then vacuum-dried overnight. Finally, the pouch was sterilized using an Anprolene AN74 ethylene oxide (EO) gas sterilizer (Andersen Sterilizers) following a standard 12-hour cycle.

[0130] The membrane topology was analyzed using a Gemini 500 scanning electron microscope (Zeiss) at the Cornell Center for Materials Research. Fiber sizes and porosity were measured with ImageJ from 10 frames taken across 3 independent preparation batches. Tensile tests were conducted using an Instron 5965 test system (Instron) and analyzed with Bluehill 3.0 SOP software.

[0131] To prepare the alginate-impregnated cell encapsulation pouch, the one-end-open fibrous pouch described above was first submerged in a 2% SLG100 alginate solution (Novamatrix) for 35 minutes to facilitate the penetration of alginate into the pores of the membrane. Subsequently, a 100 mM barium chloride solution was injected into the pouch through the open end using a 4-inch, 20-gauge blunt needle. The pouch was then held in the alginate solution for 10 seconds to allow the diffusion of barium ions across the fibrous membrane, thereby initiating gelation of the alginate both within and adjacent to the membrane. This process resulted in the formation of a thin alginate hydrogel layer that was securely affixed to the fibrous membrane. The alginate-impregnated pouch was then washed three times with normal saline and stored in a solution containing 95 mM calcium chloride and 5 mM barium chloride until further use.

[0132] Cell Loading and Assembly of BEAM System: A 4-cm section of medical-grade PTFE tubing (0.864 mm I.D. x 1.420 mm O.D.; SAI Infusion Technologies), with one end sealed 320349050v3and approximately 1.5 mm perforations along its length, was utilized as a cell loading apparatus. Pancreatic islets or INS-1 cell aggregates were washed three times with normal saline and subsequently suspended in a 2% SLG100 solution. Specifically, for the rat studies, Lewis rat pancreatic islets (6000 IEQ) were suspended in 100 µL of SLG 100 solution. The prepared cell suspension was pre-loaded into another PTFE tubing, which was then connected to the cell loading apparatus centrally positioned within the cell pouch. The other end of the PTFE tubing was connected to a syringe, which was used to gradually infuse the suspension into the cell compartment through the perforations in the loading apparatus. The outer diameter of the cell- loading device (1.420 mm) was slightly smaller than the inner diameter of the polyurethane ring in the cell encapsulation pouch (1.680 mm) to prevent pressure buildup during infusion of cell suspension. Following this, the cell encapsulation pouch was immersed in a crosslinking solution composed of 95 mM calcium chloride and 5 mM barium chloride for 10 minutes to facilitate the gelation of the cell-embedded alginate layer. The pouch was then washed 3 times with normal saline. The cell loading apparatus was removed, providing space for the subsequent insertion of oxygen transport tubing.

[0133] The oxygen transportation tubing is composed of two distinct segments: an oxygen- permeable segment constructed from USP class VI silicone tubing with an I.D. of 0.635 mm and an outer diameter O.D. of 1.194 mm (Specialty Manufacturing Inc), and an oxygen-impermeable segment fabricated from medical-grade polyurethane tubing with an I.D. of 1.016 mm and an O.D. of 1.379 mm (Scientific Commodities Inc). The length of the oxygen-permeable segment was designed to match the length of the cell-laden section of the encapsulation pouch, which was specifically 2 cm in the version used in rats. To mitigate the risk of kinking post-implantation, a miniature coil spring (D.R. Templeman Co.) was inserted into the oxygen-impermeable segment. Furthermore, a polycarbonate barb connector (Bio-Rad) was incorporated onto the oxygen- impermeable tubing. The cell encapsulation pouch was sealed by simply sliding the oxygen transportation tubing fully into the cell capsules, ensuring that the barb connector fits securely within the polyurethane ring of the cell encapsulation pouch. To assess the durability of the connection, a tensile pull test was conducted. The connector and the polyurethane ring were mounted axially in a holding fixture and tested at a rate of 15 mm per min. The force exerted at the moment of separation between the tubing and the barb was recorded. Finally, the sealed encapsulation pouch was connected to the oxygen outlet of an iEOG. All tubing connections were assembled using a custom metal barb connector (Giner Inc), and the junction was reinforced using CA-MG Infinity Bond™ cyanoacrylate adhesive (Infinity Bond). 320349050v3

[0134] Assessment of Cell Viability: The viability assessment of pancreatic islets and INS-1 cell aggregates was conducted using a dual fluorescent staining technique. Live cells were stained with acridine orange (AO; Sigma Aldrich), while dead cells were stained with ethidium homodimer-1 (EthD-1, ThermoFisher Scientific), following the provided instructions from the supplier. AO / EthD-1 were diluted in RPMI-1640 medium supplemented with 10% FBS, 10 mM HEPES, 1 mM sodium pyruvate, and 100 IU / mL penicillin and 100 μg / mL streptomycin and incubated with islets or cell aggregates for 1 hours. The fluorescent images were captured using a digital inverted fluorescent microscope (InvitrogenTMEVOSTMFL, Fisher Scientific).

[0135] Static Glucose-Stimulated Insulin Secretion (GSIS): The functionality of encapsulated pancreatic islets was assessed through a static GSIS assay. Cell encapsulation pouches from three groups—the positive control group (20% oxygen), the negative control group (1% oxygen, without oxygen supplementation), and the experimental group (1% oxygen, with oxygen supplemented from iEOGs)—were subjected to triple rinsing with Krebs-Ringer bicarbonate buffer (KRBB) and preconditioned in a 2.8 mM glucose solution for one hour at 37oC. Subsequently, these pouches were sequentially exposed to a low glucose solution (2.8 mM), followed by high glucose solutions (16.7 mM or 28 mM), and then returned to a low glucose solution (2.8 mM) under 1% O2culture condition, with each exposure period lasting one hour. Insulin concentrations in the supernatants were quantified using an enzyme-linked immunosorbent assay (ELISA) kit (Millipore Sigma). For islets encapsulated in alginate segments retrieved from animal models, the total IEQ count in the segments was determined to normalize the insulin secretion levels.

[0136] Animal Model: All animal procedures were approved by the Cornell Institutional Animal Care and Use Committee (IACUC) and supervised by the Cornell Center for Animal Resources and Education (CARE) staffs. Lewis rats (200-400 grams; Inotiv Inc) were used as pancreatic islet donors while SD rats (380-420 grams; Charles River) were used as recipients. The rats were housed within an environmentally regulated room at the Weill Hall Vivarium Animal Facility, following a standard 12-hour dark / light cycle. Daily welfare monitoring was conducted, with specific indicators including marked lethargy, hypothermia, severe dehydration, ataxia, hunched posture, labored breathing, tiptoe or slow ponderous gait, infection at the implant site, and wound dehiscence. Loss of body weight ≥ 20% compared to baseline and BCS (Body Condition Score) < 2 established as a humane endpoint. Several experiments on iEOG performance were conducted at CBSET (Lexington MA) on behalf of Giner Inc under IACUC approvals and similar daily monitoring. 320349050v3

[0137] Customized neoprene jackets were supplied by Lomir Biomedical Inc. These jackets were made with exposed flanks and an open abdominal area, a deliberate design to prevent discomfort during extended experiments. The backpack segment was seamlessly integrated into the jacket using Velcro, facilitating easy wearing and removal of the jackets. Rats were acclimated to these jackets for a period of 3 to 14 days before undergoing surgery. Rats wearing jackets were housed individually to prevent potential harm from cage mates. Objects in the cage that could entangle the jackets, such as nest boxes, were removed to avoid incidents.

[0138] Diabetes was chemically induced in SD rats by intraperitoneal administration of pharmaceutical-grade streptozotocin (STZ; Zanosar, Sicor Pharmaceutics). Initially, a freshly prepared STZ solution in 100 mM citrate buffer (pH 4.5) was injected into healthy rats (40 mg / kg), followed by a second dose (20 mg / kg) after 5 days. Blood samples were collected daily from the tail vein starting on day 7 after the first STZ dose to confirm diabetes. Rats with at least seven successive blood glucose measurements exceeding 450 mg / dL were considered diabetic and underwent the implantation procedure.

[0139] Implantation of Cell Encapsulation Pouch: To prepare for cell encapsulation pouch implantation, anesthesia was induced in rats using 4% isoflurane in oxygen and maintained at 2% throughout the surgery. The fur in the right flank was removed using Nair™ cream (Church & Dwight Company Inc), and the surgical field was ascepticized with povidone-iodine cotton swabsticks (Dynarex Corp). Then, a 2-cm skin incision was made above the latissimus dorsi muscle. A 5-cm depth subcutaneous pocket was generated parallelly with the ribs using blunt-tip scissors to make space for the implantation of the BEAM system. The cell encapsulation pouch was then introduced into the pocket. A medical-grade polyester tissue cuff (Surgical MeshTM) was integrated where the oxygen transportation tubing exited from the incision to promote healing and prevent infection. The incision was closed and secured to the transportation tubing using a cruciate suture technique. The transportation tubing was finally connected to an iEOG enclosed in a water-refillable chamber placed in the pocket secured on the back of the rat by a customized jacket. The water was replenished on a weekly basis. The overall weight of the system including internal and external components was under 10% of rat body weight. Post-Implantation Assessments

[0140] Non-Fasting Blood Glucose (NBG) Measurement: NBG levels in rats undergoing implantation were assessed at regular intervals using a Contour Next EZ blood glucose monitoring system (Bayer) following BEAM implantation. Graft failure was defined as NBG levels consistently exceeding 250 mg / dL for three consecutive days. 320349050v3

[0141] Intraperitoneal Glucose Tolerance Test (IPGTT): The metabolic function of transplanted islets in rats was evaluated through an IPGTT conducted on day 30 post- transplantation. After an 18-hour fasting period, rats received an intraperitoneal bolus injection of 50% (w / v) dextrose solution (Phoenix Pharmaceutical Inc) at a dosage of 2.0 g / kg. Blood glucose levels were measured at 0, 15, 30, 45, 60, 90, and 120 minutes following the dextrose injection. Blood samples were collected from the tail vein at 0 and 90 minutes, allowed to equilibrate at room temperature for 30 minutes, and then centrifuged at 2000 ×g for 10 minutes at 4°C to remove clotted material. The serum obtained from the supernatant was stored at -80°C. Serum C-peptide concentrations were quantified using a Rat C-peptide ELISA kit (ALPCO).

[0142] Assessment of iEOG Function on Animals: An electronic box (3.5 cm x 3.5 cm x 1.2 cm), containing a battery pack and an electronic controller, was developed by Giner Inc. to regulate the electrolysis and thereby control the oxygen production rate. The system was designed to record data on the function of the electrolyzer and battery levels at five-minute intervals. The box was equipped with a built-in infrared (IR) transmitter for communication with an IR receiver connected to a laptop, facilitating touchless monitoring of iEOG functionality from outside the animal cages. In addition, a light-emitting diode (LED) was incorporated into the system to indicate the status of oxygen production, where green indicates normal production rates and orange signals abnormal production rates. The functionality of iEOGs was assessed twice daily, and data from the electronic box were reviewed every four days to detect any malfunctions related to the electrolysis. The battery was recharged every 4-5 days to maintain continuous operation. Oxygen cessation lasting more than 24 hours was considered the endpoint of an experiment. In such cases, the cell encapsulation pouches were retrieved within 48 hours of the detected oxygen cessation.

[0143] Ex Vivo Assessment of Encapsulated Pancreatic Islets Post-Implantation: The islet- laden alginate hydrogel layer was carefully removed from the retrieved encapsulation pouch and segmented into three portions for subsequent analyses: viability assessment using AO / EthD-1 staining, functionality assessment via GSIS assay, and histological examination. For histological evaluation, the alginate segments were fixed in 10% neutral-buffered formalin solution (Sigma- Aldrich) for 24 hours. To maintain the structural integrity, the hydrogel layer was subsequently embedded in Histogel™ (Thermo Scientific). Samples were then dehydrated, embedded in paraffin (Paraplast, Leica Biosystems), and sectioned at a thickness of 5 µm using a MCT25 manual microtome (Rankin Biomedical). H&E staining was performed utilizing a commercial staining kit (Sigma) in accordance with the manufacturer's protocol. For immunofluorescence staining, tissue sections were deparaffinized and rehydrated through sequential immersion in 320349050v3xylene, followed by graded ethanol solutions (90%, 75%, 50%, 25%) and distilled water. Antigen retrieval was conducted using a pH 6.0 citrate buffer at boiling temperature, as previously described (Tran et al., “Prolongation of Graft Survival via Layer-By-Layer Assembly of Collagen and Immunosuppressive Particles on Pancreatic Islets,” Biomaterials 290:121804 (2022), which is hereby incorporated by reference in its entirety). Sections were blocked in PBS containing 1% (w / v) bovine serum albumin (BSA) and 10% (v / v) donkey serum for 2 hours at room temperature to prevent non-specific binding. Subsequently, sections were incubated overnight at 4°C with primary antibodies: rabbit anti-insulin (1:200; ab210560; Abcam) and mouse anti-glucagon (1:200; ab10988; Abcam). Following this, slides were washed five times with PBS and incubated for 1 hour at room temperature with secondary antibodies: Alexa Fluor 594-labeled donkey anti-rabbit IgG (1:400; R37119, Invitrogen) and Alexa Fluor 488-labeled donkey anti-mouse IgG (1:400; A-21202, Invitrogen). After five additional washes with PBS, the slides were mounted using Fluoroshield™ containing 4′,6-diamidino-2-phenylindole (DAPI) (Sigma). Immunofluorescence images were captured using an APEXVIEW APX100 microscope (Olympus). Statistical Analysis

[0144] All statistical analyses were performed using GraphPad Prism 10. Data are presented as individual values, mean ± standard deviation (SD), standard error of the mean (SEM), or box- and-whisker plots, as specified in the figure captions. The Shapiro-Wilk test and F-test were used to assess the normality and variance equality of the data. Statistical significance was determined at a 95% confidence level, with a p-value of less than 0.05 considered significant. For comparisons between two groups, unpaired t-tests were conducted for normally distributed data; otherwise, the Mann-Whitney U test was performed. For comparisons involving more than two groups, one-way ANOVA was utilized. All figures were generated using GraphPad Prism 10, with confidence intervals displayed for relevant data points. Example 1 – Design and Fabrication of the BEAM system

[0145] The BEAM system comprises two integral components: an implantable electrochemical oxygen generator engineered to produce a continuous and controllable supply of oxygen through the electrolytic conversion of water and a cell-encapsulating pouch made of alginate-impregnated electrospun fibrous membrane for immunoprotection. Given biocompatibility is a pivotal requisite for the clinical translation of biomedical devices, all the components of the BEAM system were engineered from medical-grade materials. iEOGs were GMP-manufactured by Giner Inc. The electrolyzer components were housed within a medical- 320349050v3grade titanium enclosure with a porous polymeric membrane integrated onto one face, functioning as a window to harvest interstitial water vapor from surrounding tissue for electrolysis. The iEOGs were controlled to a settable current with corresponding voltages from 1.4 to 1.8 V, in the range for the splitting of water into hydrogen and oxygen, which were released through two separate outlets (FIG.1A). The oxygen outlet delivered gas by convection to an oxygen-permeable tubing traversing the cylindrical cell encapsulation pouch, enabling oxygen diffusion along its length to the encapsulated cells (FIGs.1B-1C). An alginate- impregnated fibrous membrane was used as the immuno-protective barrier, which allows for selective diffusion of nutrients, glucose, and insulin while preventing the infiltration of host cells.

[0146] Two generations of the iEOG were developed by Giner Inc based on the membrane electrode assembly (MEA) technology as previously described (U.S. Patent No.11,773,496 to Schwenk et al.; U.S. Patent No.10,231,817 to Tempelman et al., which are hereby incorporated by reference in their entirety) (FIG.2A). The MEA consists of a proton exchange membrane that when hydrated serves as the electrolyte, transferring the protons in the balanced electrochemical reaction. The anode and cathode electrocatalysts are pressed on opposite sides of the membrane as a single piece composite. At the surface of the anode, oxygen is evolved and leaves via convection through the oxygen gas port on the anode side. At the surface of the cathode, hydrogen is evolved and leaves via convection through the hydrogen gas port on the cathode side. The first version (v1 iEOG) had dimensions of 20 mm in diameter, 4.6 mm in height, and a weight of approximately 4 grams. The second version (v2 iEOG) was designed to be smaller in both diameter and thickness by decreasing the MEA area, while maintaining an equivalent oxygen generation capacity. Specifically, the v2 iEOG measures 13 mm in diameter, 3.1 mm in height, and approximately 2 grams in weight, which is approximately 25% of the volume and 50% of the weight of the v1 iEOG. Both versions demonstrated precise regulation of oxygen production via modulation of electrical current, allowing for variable oxygen generation rates tailored to meet the metabolic demands of different cells and cell doses, ranging from small numbers (~1000 IEQ) to clinically relevant doses of islets.

[0147] The average dose of cadaveric pancreatic islets transplanted into the portal vein during the U.S. CIT-07 trials (NCT00434811) was approximately 800,000 IEQ (Ricordi et al., “National Institutes of Health-Sponsored Clinical Islet Transplantation Consortium Phase 3 Trial: Manufacture of a Complex Cellular Product at Eight Processing Facilities,” Diabetes 65(11):3418-3428 (2016), which is hereby incorporated by reference in its entirety). While the optimal dose of islets for extrahepatic transplantation sites has not yet been established, it was 320349050v3predicted that the required dose would range from 300,000 IEQ to 770,000 IEQ for a 70 kg patient (Owyang et al., “Re-Considering Quantity Requirements in Islet Transplantation,” Nature Reviews Bioengineering 1(6):382-384 (2023), which is hereby incorporated by reference in its entirety). To evaluate whether the oxygen generation capability of the system could meet the oxygen demand of clinically relevant doses of islets over extended periods, three v1 iEOGs were operated at 10 mA in saline for 11 months, 2 years, and 2.5 years. All three devices exhibited consistent currents close to the setpoint throughout the duration of the study, resulting in a stable oxygen output of approximately 1.94 standard cubic centimeters per hour (scch) (FIG.2B). This oxygen flow rate is sufficient to support >600,000 human IEQ, assuming an oxygen consumption rate of 2.1 pmol / min / IEQ (Table 1). To assess whether the iEOGs could achieve similar oxygen production in vivo, the devices were implanted in immunodeficient nude rats or immunocompetent Sprague-Dawley (SD) rats, operating at 11 mA to simulate expected clinical conditions. After three months of implantation in RNU nude rats, the v1 iEOG generated comparable oxygen levels (2.31 ± 0.10 scch) to those measured pre-implantation (2.25 ± 0.01 scch), which were calculated to be sufficient to support 720,000-740,000 human IEQ (FIG.2C). Similar results were observed for the v2 iEOG, with oxygen production levels of 2.29 ± 0.03 scch maintained after two months of implantation in nude rats (FIG.2D). In immunocompetent SD rats, the v2 iEOG also demonstrated stable oxygen generation, with levels measured at 2.44 ± 0.16 scch before implantation and 2.29 ± 0.12 scch one month post-implantation (FIG.2E). Taken together, these findings suggest the long-term capability of iEOGs to produce sufficient oxygen from interstitial moisture to support clinically relevant doses of human pancreatic islets. Furthermore, histological analysis using Masson’s trichrome staining of tissue surrounding the iEOG in SD rats after one month showed no evidence of adverse tissue reactions and indicated the presence of blood vessels near the water-harvesting window (FIGs.2F-2G). The v2 iEOG was selected for further development given the compact design and reduced weight, making it more suitable for implantation. 320349050v3Table 1. Oxygen Consumption Rates of Insulin-Secreting CellsEach of references (1) and (3)-(13), listed below, is hereby incorporated by reference in its entirety: (1) Ernst et al., “A Predictive Computational Platform for Optimizing the Design of Bioartificial Pancreas Devices,” Nature Communications 13:6031 (2022); (3) Cline et al., “Rates of Insulin Secretion in INS-1 Cells are Enhanced by Coupling to Anaplerosis and Kreb's Cycle Flux Independent of ATP Synthesis,” Biochem Biophys Res Commun 415:30-35 (2011); (4) Dover et al., “Arsenite and Methylarsonite Inhibit Mitochondrial Metabolism and Glucose-Stimulated Insulin 320349050v3Secretion in INS-1832 / 13 β cells,” Arch Toxicol 92:693-704 (2018); (5) Rumala et al., “Exposure of Pancreatic β-Cells to Excess Glucose Results in Bimodal Activation of mTORC1 and mTOR-Dependent Metabolic Acceleration,” iScience 23:100858 (2020); (6) Evron et al., “Long-Term Viability and Function of Transplanted Islets Macroencapsulated at High Density are Achieved by Enhanced Oxygen Supply,” Scientific Reports 8:6508 (2018); (7) Papas et al., “A Stirred Microchamber for Oxygen Consumption Rate Measurements with Pancreatic Islets,” Biotechnol Bioeng 98:1071-1082 (2007); (8) Pisania, “Development of Quantitative Methods for Quality Assessment of Islets of Langerhans,” Thesis, Massachusetts Institute of Technology (2007); (9) Suszynski et al., “Islet Size Index as a Predictor of Outcomes in clinical Islet Autotransplantation,” Transplantation 97:1286-1291 (2014); (10) Papas et al., “Human Islet Oxygen Consumption Rate and DNA Measurements Predict Diabetes Reversal in Nude Mice,” American Journal of Transplantation 7:707-713 (2007); (11) Balboa et al., “Functional, Metabolic and Transcriptional Maturation of Human Pancreatic Islets Derived From Stem Cells,” Nature Biotechnology 40:1042-1055 (2022); (12) Papas et al., “Islet Oxygen Consumption Rate (OCR) Dose Predicts Insulin Independence in Clinical Islet Autotransplantation,” PLoS One 10:e0134428 (2015); and (13) Wang et al., “Increased Oxygen Consumption Rates in Response to High Glucose Detected by a Novel Oxygen Biosensor System in Non-Human Primate and Human Islets,” J Endocrinol 185:445-455 (2005). For ease of comparison across studies employing varying units, the original values are presented together with the estimated values in alternative units (marked with (*)). The conversion is based on the assumption that 1 IEQ corresponds to 1,560 cells, containing 10.4 ng of DNA.

[0148] The cell encapsulation pouch was designed to be mechanically robust, scalable, and biocompatible. Mechanical properties, specifically stiffness and elasticity, significantly impact the foreign body responses following implantation (Capuani et al., “Advanced Strategies to Thwart Foreign Body Response to Implantable Devices,” Bioeng Transl Med 7(3):e10300 (2022); Kämmerling et al., “Mitigating the Foreign Body Response Through ‘Immune- Instructive’ Biomaterials,” Journal of Immunology and Regenerative Medicine 12:100040 (2021); Ni et al., “Macrophages Modulate Stiffness-Related Foreign Body Responses Through Plasma Membrane Deformation,” Proceedings of the National Academy of Sciences 120(3):e2213837120 (2023); Carnicer-Lombarte et al., “Foreign Body Reaction to Implanted Biomaterials and its Impact in Nerve Neuroprosthetics,” Frontiers in Bioengineering and Biotechnology 9:622524 (2021), which are hereby incorporated by reference in their entirety), which is a crucial determinant for the outcomes of a cell encapsulation implant. Ideally, the pouch should be soft to reduce the mechanical mismatch with the surrounding host tissue, which can trigger the activation of inflammatory and fibroblastic cells (Noskovicova et al., “Suppression of the Fibrotic Encapsulation of Silicone Implants by Inhibiting the Mechanical Activation of Pro-Fibrotic TGF-β,” Nature Biomedical Engineering 5(12):1437-1456 (2021); Rolfe et al., “The Fibrotic Response to Implanted Biomaterials: Implications for Tissue 320349050v3Engineering,” in Regenerative Medicine and Tissue Engineering, IntechOpen, London (2011); Klopfleisch et al., “The Pathology of the Foreign Body Reaction Against Biomaterials,” J Biomed Mater Res A 105(3):927-940 (2017), which are hereby incorporated by reference in their entirety). On the other hand, it also needs to be elastic and durable to maintain its structural integrity during surgical handling and long-term indwelling as intensified fibrotic responses were generally observed in the rough and kinked regions caused by implant deformation (Thanos et al., “Considerations for Successful Encapsulated β-Cell Therapy, in Cell Therapy: Current Status and Future Directions,” D.F. Emerich and G. Orive, Eds., Springer International Publishing: Cham. p.19-52 (2017), which is hereby incorporated by reference in its entirety) (FIG.3). To meet these requirements, an electrospinning technique was used to prepare a soft, highly porous pouch with round end using medical grade polyether block amide (PEBAX) as a base material (FIG.4A). Arkema Pebax® thermoplastic elastomer was selected for its inherent softness (77 Shore D) and flexible, rubber-like nature (Flexural modulus of 12). Scanning electron microscopy (SEM) of the membrane revealed a fibrous structure with interconnected pores, which is essential for efficient mass transfer (Figure 4B-4C). The fibers were randomly aligned with sizes ranging from 1.4 to 4 µm (average size: 2.47 ± 0.43 µm) (FIG.4D); this relatively large fiber sizes were chosen to facilitate mass transfer and improve capillary-driven penetration of alginate into the membrane during the hydrogel coating process. The porosity of the membrane was estimated to be 55.12 ± 5.93% (FIG.5). The thickness of the membrane was approximately 70 µm, as determined by SEM. Furthermore, the membrane demonstrated considerable mechanical robustness and elasticity, as indicated by a tensile strength greater than 8 MPa and a fracture strain exceeding 23 (FIG.4E).

[0149] One of the challenges in the fabrication of membrane-based encapsulation systems is achieving effective sealing. Conventional sealing methods involving the use of heat, ultrasonic welding, or adhesives can be troublesome due to interference from liquid (media, crosslinking solution) used during the cell loading process. In addition, these methods generally result in rigid and sharp edges, which potentially exacerbate the fibrotic responses. The by-products generated during the application of heat or glue may further contribute to these adverse reactions. Indeed, significantly enhanced fibrotic responses were observed at the heat-sealed ends of membrane- based systems compared to the membranes themselves, regardless of the membranes' physicochemical properties, across various immunocompetent animal models, including C57BL / 6 mice (FIGs.6-9), SD rats, and Göttingen minipigs (Liu et al., “A Zwitterionic Polyurethane Nanoporous Device with Low Foreign-Body Response for Islet Encapsulation,” Adv Mater 33(39):e2102852 (2021), which is hereby incorporated by reference in its entirety). 320349050v3The necessity of employing heat sealers, welders, or adhesives also complicates the fabrication process, especially under aseptic conditions. To address these challenges, an edge-free cell pouch with a specific geometry was designed to minimize the area requiring sealing (FIG.4A). A water-soluble polyethylene glycol (PEG) template was employed to collect the electrospun fibers, thus shaping the cell encapsulation compartment into an edge-free round-ended pouch (FIG.10). A medical-grade polyurethane ring was incorporated into the other open end to serve as a female component in a barbed slide-in connection. This configuration allows the pouch to be sealed by simply sliding the oxygen transportation tubing into the lumen of the cell encapsulation pouch.

[0150] FIG.4B shows that the encapsulation pouch can be scaled both radially and longitudinally to accommodate different doses of cells. For long pouches (e.g. human version) which tend to kink, the membrane was reinforced with an elastic, shape-memorable nitinol braided mesh scaffold with a wire thickness of 25.4 µm and pore sizes ranging from 100 µm to 150 µm (FIGs.4F and 11). The reinforced pouches did not kink and were able to revert to their original form after common deformations, such as folding, bending, or twisting, that may occur during the surgical procedure or host movement (FIG.4G). The impact of the nitinol scaffold on mass transfer is expected to be negligible owning to the large pore sizes and thin wire thickness. To impart the immuno-protective property, the fibrous membrane was impregnated with ultrapure alginate (SLG100) using a previously developed method to avoid the detachment of alginate hydrogel after implantation (Liu et al., “A Safe, Fibrosis-Mitigating, and Scalable Encapsulation Device Supports Long-Term Function of Insulin-Producing Cells,” Small 18(8):e2104899 (2022), which is hereby incorporated by reference in its entirety). This resulted in a hydrogel layer thickness of 30-50 µm, bringing the final thickness of the pouch wall to approximately 100-120 µm (FIG.4H).

[0151] A customized cell-loading apparatus, constructed from a perforated polytetrafluoroethylene (PTFE) tube, was developed for cell loading into the BEAM system. PTFE was chosen due to its inertness and hydrophobic properties, which effectively reduce the adhesion of residual alginate solution, thereby minimizing cell loss during encapsulation. To demonstrate the loading procedure, INS-1 cell aggregates with a size distribution from 50 to 300 μm were used as a model to mimic pancreatic islets (FIG.12). These aggregates were gently dispersed in a 2% alginate solution (SLG100) at a density of 60,000 IEQ / mL before being introduced into the cell-loading apparatus. FIG.13A shows the streamlined procedure of cell loading and system sealing. To better visualize the cell loading process, 150-µm beads (used to simulate cell aggregates) were loaded through the loading apparatus, into an encapsulation pouch 320349050v3made of a transparent tube, resulting in a relatively even distribution within the annular space between the loading tool and the pouch wall (FIG.14). When the encapsulation pouch was immersed in a crosslinking solution containing 95 mM calcium chloride and 5 mM barium chloride, the divalent ions diffused through the porous membrane, initiating alginate gelation. Both longitudinal and cross-sectional histology images of the pouches revealed the annular configuration of the aggregate-laden alginate layer with the thickness of 300-500 µm, which is equivalent to about 2-3 layers of cell aggregates (FIG.13B). This thin hydrogel layer was found to adhere to the fibrous membrane, thus reducing the distance between the cells and the pouch surface to enhance mass transfer. Live / dead staining and histological examination showed no significant changes in aggregate morphology or viability after loading into the BEAM system (FIGs.13B-13C).

[0152] Upon crosslinking of the alginate layer, the loading apparatus was withdrawn, allowing for the insertion of the oxygen-transportation tubing. This tubing comprised end-sealed silicone tubing connected to gas-impermeable polyurethane tubing, thereby facilitating oxygen diffusion selectively through the silicone wall to support adjacent cells (FIG.13D). A barbed connector was integrated onto the gas-impermeable section, designed to fit with the polyurethane ring of the cell encapsulation pouch. Once the oxygen-transportation tubing was fully inserted, the barbed connector provided a tight fit with the ring, effectively sealing the pouch to prevent gas leakage and maintain immunoprotection (FIG.13E). The connection demonstrated substantial robustness, with the ability to withstand pull forces exceeding 8.5 N (FIG.13F). The sealed cell encapsulation pouch was finally connected to an iEOG using a custom barb connector before implantation. Example 2 – Oxygenation by the BEAM System Preserved the Viability and Function of INS-1 Aggregates Under an in vitro Hypoxic Condition

[0153] The local partial oxygen tension surrounding an encapsulation implant is a critical determinant for the survival and function of encapsulated β-cells, particularly when these cells are loaded at a high density. Unfortunately, post-transplantation oxygen levels in the vicinity of subcutaneous implants have been reported to be critically low (<10 mmHg), primarily due to inadequate vascularization at the subcutaneous implantation site and the subsequent formation of a fibrotic capsule (Einstein et al., “Hypoxia Within Subcutaneously Implanted Macroencapsulation Devices Limits the Viability and Functionality of Densely Loaded Islets,” Front Transplant 2:1257029 (2023); Najdahmadi et al., “Non-Invasive Monitoring of Oxygen Tension and Oxygen Transport Inside Subcutaneous Devices After H2S Treatment,” Cell 320349050v3Transplantation 29:0963689719893936 (2020); Mitsugashira et al., “Development of a Novel Method for Measuring Tissue Oxygen Pressure to Improve the Hypoxic Condition in Subcutaneous Islet Transplantation,” Scientific Reports 12(1):14731 (2022), which are hereby incorporated by reference in their entirety). Given the inherently high oxygen consumption rate of β-cells, such low oxygen tension is insufficient to support their long-term survival and function. In addition, this also limits the feasibility of loading them at a high density, thus rendering the requisite implant size impractical for clinical applications. It was assumed that enhancing the oxygen supply to the implant could improve cell survival and elevate the upper limit of islet cell density. To test this hypothesis, INS-1832 / 13 aggregates (2000 IEQ), an insulin-secreting cell line with oxygen consumption rates comparable to that of rat pancreatic islets and higher than that of human islets (Table 1), were encapsulated in a BEAM system, resulting in a relatively high loading density (60,000 IEQ / mL). These implants were challenged under 1% O2cell culture condition (~ 7.62 mmHg) for 24 hours (FIGs.15A-15B). Significant cell death was observed in non-oxygenated implants, as evidenced by cell disintegration, shrunken morphology, and live / dead staining (FIGs.15C-15D). Histological analysis further revealed a fragmented aggregate structure and cell apoptosis evidenced by weak cytoplasmic staining, nuclear shrinkage, and the formation of apoptotic bodies in the majority of cells (FIG. 15E). Additionally, insulin-positive cells were scarcely detected in these non-oxygenated systems (FIG.15F). These findings demonstrate a substantial reduction in the viability and function of INS-1832 / 13 cells following 24 hours of culture under a hypoxic condition. In contrast, cell aggregates in systems supplied with an oxygen generated at an electrical current of 270 µA maintained normal morphology and healthy aggregate structure, with a negligible dead cell number, as indicated by both live / dead staining and histological examination (FIGs.15B- 15F). Cell aggregates in regions adjacent to silicone tubing (proximal to the oxygen source) and regions near the fibrous membrane (distal to oxygen source) remained highly viable (FIG.16). The oxygen generation rate was approximately 0.016 scch, as estimated using the simulation. In addition, the majority of cells in this group were positive for insulin staining, suggesting that oxygenation by the BEAM system effectively preserved INS-1832 / 13 cell aggregate function under a hypoxic culture condition, even at a relatively high cell density. Example 2 – Oxygenation by the BEAM System Preserved the Viability and Function of Human Islets Under an in vitro Hypoxic Condition

[0154] The impact of oxygenation provided by the BEAM system on the viability and function of primary human pancreatic islets was then assessed. Approximately 1,800 human IEQ 320349050v3were dispersed in 30 µL of 2% SLG100 solution and subsequently loaded into a BEAM system at a density of 60,000 IEQ / mL and subjected to 1% O2culture condition (FIGs.17A-17B). The BEAM system was operated at 240 µA, corresponding to an oxygen generation rate of approximately 0.009 scch, as estimated by our simulation. After 24 hours, pancreatic islets encapsulated in systems without oxygenation exhibited significant cell dissociation and apoptosis (FIGs.17C-17D). Immunofluorescence staining revealed a notable decline in cell functionality, as indicated by dominant loss in insulin and glucagon expression compared to those of healthy islets (FIG.17E). The impaired function was further corroborated by glucose stimulated insulin section (GSIS) assays (FIG.17F). The levels of insulin secreted from systems cultured under the normoxic condition for 24 hours measured 405.6 ± 16.6 µIU / h and 745.9 ± 228.0 µIU / h in the 2.8 mM and 28 mM glucose solutions, respectively. Meanwhile, non- oxygenated systems exhibited significantly lower insulin secretion levels, with 25.5 ± 4.8 µIU / h in a 2.8 mM glucose solution and 99.1 ± 80.9 µIU / h in a 28 mM glucose solution. In contrast, live / dead staining and histological analysis demonstrated that pancreatic islets encapsulated in systems supplied with oxygen remained viable after 72 hours of incubation under hypoxic culture condition (FIGs.17C-17D). A minor percentage of dead islets was observed at the edges of alginate slabs, likely due to mechanical disruption during sectioning of the hydrogel layer. GSIS and histological evaluations indicated that the function of these islets was comparable to those incubated under normal culture conditions. After 24 hours of hypoxic incubation, insulin secretion levels from these oxygenated systems were 330.7 ± 92.0 µIU / h in the 2.8 mM glucose solution and 1,025.5 ± 172.2 µIU / h in the 28 mM glucose solution. Notably, strong insulin and glucagon staining were observed in pancreatic islets of this group after 24 hours of incubation (FIG.17E). These findings suggest that the continuous supply of oxygen by the BEAM system effectively preserved the viability and function of human pancreatic islets encapsulated at a density of 60,000 IEQ / mL. Example 4 – Effects of Oxygenation by the BEAM System in an Allogeneic Rat Transplantation Model

[0155] After validating the favorable effect of oxygenation in vitro, the investigation was extended to assess its potential to enhance the viability and function of pancreatic islets in vivo using an allogeneic rat model. Pancreatic islets (6000 IEQ) isolated from Lewis rats were encapsulated in a cell pouch (3.2 mm O.D. x 2 cm length; loading capacity of ~100 µL) of the BEAM system and transplanted into STZ-induced diabetic, immunocompetent SD rats. All rats 320349050v3selected for implantation exhibited blood glucose levels exceeding 450 mg / dL for over 7 consecutive days. The weights of rats before implantation were 418.8 ± 58.5 grams, resulting in the islet doses of 14,509.5 ± 1,729.6 IEQ / kg. Owing to the relatively large size of the current electronic controller (3.5 cm x 3.5 cm x 1.2 cm) designed for bench testing, it was not feasible to fully implant the entire BEAM system within the subcutaneous space of rats. Thus, we used an extracorporeal setting, in which electronic components including battery, electronic controller, and iEOG enclosed in a refillable water reservoir, were secured in a pocket worn by the rats (FIGs.18A-18B and 19). This setting enables convenient monitoring of iEOG function, water consumption, and battery exchange. For instance, the extracorporeal electronic box allows for the incorporation of a light-emitting diode (LED) to indicate the status of oxygen production for early detection of electrolyzer malfunction or system disconnections, which is crucial in long- term animal study. It is worth noting that for clinical applications, the entire system would be implanted with rechargeable power supply charged weekly via transcutaneous energy transfer (FIG.20). For the rat study, a customized animal jacket with a small pocket on the back was designed to secure these electronic components. The flank and abdominal area of the jacket were left open to minimize the discomfort or stress on the rats (FIG.21). The use of jackets did not significantly impair the mobility of rats, nor did it cause negative health issues, as evidenced by steady body weight gain over a month (FIG.22). The battery was exchanged every 5 days, and the water reservoir was replenished on a weekly basis. The iEOGs were operated at 280 µA to achieve oxygen generation rates of 0.455 ± 0.015 scc per day (FIGs.18C-18D). The cell encapsulation pouch was implanted subcutaneously through a minor skin incision and connected to the iEOG via percutaneous oxygen-impermeable polyurethane tubing. A medical-grade polyester tissue cuff was incorporated at the incision to aid healing and prevent infection. In all 9 animals that underwent transcutaneous implantation, complete wound closure was observed without notable signs of inflammation or adverse reactions at the incision site (FIGs.23A-23B). Implants without connection to an iEOG (4 / 4) failed to reverse diabetes, whereas all rats (9 / 9) that received an iEOG-connected implant returned to normoglycemia within 3 days (FIGs.18E- 18G). Early oxygenation cessation occurred in 3 rats from the first cohort on day 1, day 3, and day 5 because of the damage of the transcutaneous tubing and electronic wire, which were exposed out of the jackets and compromised by the animals (FIG.24; Table 2). The implantation location and jacket were then adjusted so these components were secured under the jacket. As a result, the remaining rats with continuous oxygenation maintained normoglycemia for over a month and up to 88 days. Intraperitoneal glucose tolerance test (IPGTT) on day 35 indicated that these rats tolerated glucose comparably to healthy rats, suggesting the normal metabolic function 320349050v3of encapsulated islets (FIGs.18H-18I and 25) without significant mass transport limitations. This was further corroborated by the significant increases in serum C-peptide after glucose administration (FIG.26) and the body weight differences between the oxygenated and non- oxygenated groups (FIG.18G). In contrast, the blood glucose levels of rats implanted with non- oxygenated implants remained higher than 400 mg / dL after 120 min. Oxygenation was accidentally stopped in 2 rats on day 51 and day 72 due to the disconnection of the iEOG with the electronic controller and the cell pouch caused by the rats (FIG.24; Table 2). Intriguingly, sharp increases in blood glucose levels were observed right after oxygenation cessation at both early time points (day 1, 3, and 5) and later time points (day 51 and day 72). This indicates that a continuous supply of oxygen may be vital for the long-term function of pancreatic islets in cell encapsulation systems, particularly at high loading density. This is in contrast to a hypothesis in the encapsulation field that short-term oxygenation in the early period may be sufficient. Table 2. Summary of Incidents That Occurred During the Animal Study

[0156] Post-retrieval evaluations further revealed prominent cell death with shrunken morphology in implants without oxygenation and implants with ceased oxygenation, as indicated by both live / dead staining and histology examination (FIG.27A-27D and 28-29). Almost no insulin-positive cells were found in these implants (FIG.27C). No signs of host cell infiltration were observed in these systems, suggesting the destruction of pancreatic islets was not likely caused by immune cell-mediated rejection. GSIS assay demonstrated low and no significant differences between insulin levels secreted by these cells under a low glucose condition (2.8 mM) and a high glucose condition (16.7 mM), suggesting compromised function of β-cells (FIG. 320349050v327E). In contrast, viable islets with healthy morphology and strong insulin / glucagon staining were found in the oxygenated implant on day 32, 40, 45, and 88, suggesting the feasibility of oxygenation to preserve viability and function. GSIS assay on the islets retrieved from these implants showed substantial levels of secreted insulin and the ability to respond to glucose level change. It is worth noting that among these implants, three were retrieved after the blood glucose levels started to increase. A gas bubble partially surrounding the implants was observed in the rats with escalated blood glucose levels (3 / 6), probably due to the accumulation of excessive oxygen that separated the implants from the surrounding tissue (FIGs.30A-30E). To confirm this hypothesis, excessive gas was vented through a 23G needle puncture in a rat, quickly restoring normal blood glucose levels within a few hours and maintaining normoglycemia for an additional 2 weeks until oxygenation accidentally stopped on day 72. Discussion of Examples 1-4

[0157] Macroencapsulation remains a compelling strategy for cell therapy in the treatment of T1D as the use of a permselective membrane could potentially obviate the need for chronic immunosuppression. An increasing body of successful proof-of-concept preclinical studies in mice has demonstrated the potential of this approach to protect encapsulated insulin-secreting cells from immune rejection, thereby enabling long-term glucose homeostasis without the need for immunosuppression (An et al., “Designing a Retrievable and Scalable Cell Encapsulation Device for Potential Treatment of Type 1 Diabetes,” Proceedings of the National Academy of Sciences 115(2):E263-E272 (2018); Bose et al., “A Retrievable Implant for the Long-Term Encapsulation and Survival of Therapeutic Xenogeneic Cells,” Nature Biomedical Engineering 4(8):814-826 (2020); Grattoni et al., “Harnessing Cellular Therapeutics for Type 1 Diabetes Mellitus: Progress, Challenges, and the Road Ahead,” Nature Reviews Endocrinology 21:14–30 (2025); Chendke et al., “Replenishable Prevascularized Cell Encapsulation Devices Increase Graft Survival and Function in the Subcutaneous Space,” Bioeng Transl Med 8(4):e10520 (2023); Wang et al., “A Nanofibrous Encapsulation Device for Safe Delivery of Insulin- Producing Cells to Treat Type 1 Diabetes,” Science Translational Medicine 13(596):eabb4601 (2021); Nyitray et al., “Polycaprolactone Thin-Film Micro- and Nanoporous Cell-Encapsulation Devices,” ACS Nano 9(6):5675-5682 (2015); Veiseh et al., “Size- and Shape-Dependent Foreign Body Immune Response to Materials Implanted in Rodents and Non-Human Primates,” Nature Materials 14(6):643-651 (2015); Vegas et al., “Combinatorial Hydrogel Library Enables Identification of Materials that Mitigate the Foreign Body Response in Primates,” Nature Biotechnology 34(3):345-352 (2016); Mukherjee et al., “Screening Hydrogels for Antifibrotic 320349050v3Properties by Implanting Cellularly Barcoded Alginates in Mice and a Non-Human Primate,” Nature Biomedical Engineering 7(7):867-886 (2023), which are hereby incorporated by reference in their entirety). However, the translation of these findings to larger animal models, as well as the scaling up of implants for clinical applications, remains a significant challenge. While a few hundred pancreatic islets, loaded within reasonably sized implants at low densities, may be sufficient to reverse diabetes and maintain normoglycemia in mouse models, scaling up these implants for use in larger animals and eventually humans is considerably more complex. The complexity arises from the necessity to pack islets at a high density to achieve an implant size that is practical for implantation. Pancreatic islets have a well-documented reliance on oxygen for their insulin secretion in response to glucose changes. Beta cells can survive under partial oxygen tension levels exceeding 0.1-0.44 mmHg while levels greater than 10 mmHg are necessary to sustain their ability for insulin secretion (Colton, C.K., “Oxygen Supply to Encapsulated Therapeutic Cells,” Advanced Drug Delivery Reviews 67-68:93-110 (2014); Papas et al., “Effect of Oxygen Supply on the Size of Implantable Islet-Containing Encapsulation Devices,” Panminerva Med 58(1):72-7 (2016), which are hereby incorporated by reference in their entirety). Unfortunately, the local oxygen tension within encapsulation systems is significantly below the threshold required for the full function of pancreatic islets when transplanted into the poorly vascularized subcutaneous space (Einstein et al., “Hypoxia Within Subcutaneously Implanted Macroencapsulation Devices Limits the Viability and Functionality of Densely Loaded Islets,” Front Transplant 2:1257029 (2023); Najdahmadi et al., “Non-Invasive Monitoring of Oxygen Tension and Oxygen Transport Inside Subcutaneous Devices After H2S Treatment,” Cell Transplantation 29:0963689719893936 (2020); Mitsugashira et al., “Development of a Novel Method for Measuring Tissue Oxygen Pressure to Improve the Hypoxic Condition in Subcutaneous Islet Transplantation,” Scientific Reports 12(1):14731 (2022), which are hereby incorporated by reference in their entirety). This is further exacerbated by the additional mass transport barrier caused by the membrane and the subsequent formation of a fibrotic capsule, which is known to be more severe in larger animals and humans. Results from clinical trials by Viacyte Inc with an immunoprotective device (PEC-Encap™) and an open device (PEC-Direct™) loaded with pancreatic endoderm cells suggest that even with the use of membranes designed to promote vascularization or the membrane with perforations, the devices may fail to achieve sufficient vascularization in both timing and density to support a metabolically adequate functional beta cell mass (Keymeulen et al., “Encapsulated Stem Cell– Derived β Cells Exert Glucose Control in Patients with Type 1 Diabetes,” Nature Biotechnology 42(10):1507-1514 (2024), which is hereby incorporated by reference in its entirety). Moreover, 320349050v3increasing cell density further heightens oxygen demand due to the steeper oxygen gradients within cell layers. Thus, in the previously developed cell encapsulation systems, insulin- secreting cells were typically loaded at densities not exceeding 25,000 IEQ / mL (Table 3). This would necessitate impractically large encapsulation systems to accommodate a clinically relevant dose of pancreatic islets (300,000 IEQ-770,000 IEQ) at such low densities. For instance, in the clinical trial by ViaCyte (NCT04678557), 12 planar devices with a surface area of approximately 10 cm2for each were required to achieve a metabolically relevant dose of cells. Computational modeling simulations suggest that a fiber measuring 12.8 meters in length with a 1 mm diameter, or a circular slab with a diameter of 11.9 cm and a thickness of 0.5 mm, would be required to deliver 500,000 functional IEQ under 40 mmHg O₂ (Ernst et al., “A Predictive Computational Platform for Optimizing the Design of Bioartificial Pancreas Devices,” Nature Communications 13(1):6031 (2022), which is hereby incorporated by reference in its entirety). Even with microencapsulation, which provides a higher surface-to-volume ratio as compared to macroencapsulation systems, rapid decline in functionality was observed when the loading density was increased to 16,000 IEQ / mL (Mukherjee et al., “Screening Hydrogels for Antifibrotic Properties by Implanting Cellularly Barcoded Alginates in Mice and a Non-Human Primate,” Nature Biomedical Engineering 7(7):867-886 (2023), which is hereby incorporated by reference in its entirety). Thus, oxygen supplementation is likely needed to realize the clinical translation of cell encapsulation. Table 3. Loading Densities Employed in Micro- and Macro-Encapsulation Systems Type of encapsulation Loading density Cell type Ref.Microencapsulation 4000-16,000 IEQ / mL Human islets (17) Microencapsulation 2000 cell clusters / mL SC-beta cells (18) Microencapsulation 4000 IEQ / mL Porcine islets (19) oxygenationMicroencapsulation 200-2000 cell clusters / mL SC-beta cells (20)Macroencapsulation 25,000 IEQ / mL Rat islets (21) Macroencapsulation 8300 IEQ / mL Rat and human islets Macroencapsulation 25,000 islets / mL Rat isletsMacroencapsulation 25,000 IEQ / mL Rat islets (24) WithMacroencapsulation 2250 IEQ / cm2 Rat islets (25)oxygenationMacroencapsulation 1000-4800 IEQ / cm2 Rat islets (26)Macroencapsulation 6250-8330 IEQ / mL Rat islets (27) Macroencapsulation1000 IEQ / cm2or 21,250Rat islets (28) islets / mL Macroencapsulation 60,000 IEQ / mL or 4200 Rat islets Present IEQ / cm2disclosure Each of references (14)-(28), listed below, is hereby incorporated by reference in its entirety: 320349050v3(14) Hu et al., “Toll-Like Receptor 2-Modulating Pectin-Polymers in Alginate-Based Microcapsules Attenuate Immune Responses and Support Islet-Xenograft Survival,” Biomaterials 266: 120460 (2021); (15) Qin et al., “Pancreatic Stellate Cells Support Human Pancreatic β-Cell Viability in Vitro and Enhance Survival of Immunoisolated human Islets Exposed to Cytokines,” Materials Today Bio 27:101129 (2024); (16) Veiseh et al., “Size- and Shape-Dependent Foreign Body immune Response to Materials Implanted in Rodents and Non-Human Primates,” Nature Materials 14:643-651 (2015); (17) Mukherjee et al., “Screening Hydrogels for Antifibrotic Properties by Implanting Cellularly Barcoded Alginates in Mice and a Non-Human Primate,” Nat Biomed Eng 7:867-886 (2023); (18) Alagpulinsa et al., “Alginate-Microencapsulation of Human Stem Cell–Derived β Cells With CXCL12 Prolongs Their Survival and Function in Immunocompetent Mice Without Systemic Immunosuppression,” American Journal of Transplantation 19:1930-1940 (2019); (19) Holdcraft et al., “Enhancement of in Vitro and in Vivo Function of Agarose-Encapsulated Porcine Islets by Changes in the Islet Microenvironment,” Cell Transplantation 23:929-944 (2014), (20) Vegas et al., “Long-Term Glycemic Control Using Polymer-Encapsulated Human Stem Cell–Derived Beta Cells in Immune-Competent Mice,” Nature Medicine 22:306-311 (2016); (21) Bose et al., “A Retrievable Implant for the Long-Term Encapsulation and Survival of Therapeutic Xenogeneic Cells,” Nat Biomed Eng 4:814-826 (2020); (22) An et al., “Designing a Retrievable and Scalable Cell Encapsulation Device for Potential Treatment of Type 1 Diabetes,” Proceedings of the National Academy of Sciences 115:E263-E272 (2018); (23) Jesser et al., “Pancreatic Islet Macroencapsulation: a New Device for the Evaluation of Artificial Membrane,” Artif Organs 20:997-1007 (1996); (24) Kumagai-Braesch et al., “The TheraCyte™ Device Protects Against Islet Allograft Rejection in Immunized Hosts,” Cell Transplant 22:1137-1146 (2013); (25) Barkai et al., “Enhanced Oxygen Supply Improves Islet Viability in a New Bioartificial Pancreas,” Cell Transplant 22:1463-1476 (2013); (26) Evron et al., “Long-Term Viability and Function of Transplanted Islets Macroencapsulated at High Density are Achieved by Enhanced Oxygen Supply,” Sci Rep 8:6508 (2018); (27) Wang et al., “An Inverse-Breathing Encapsulation System for Cell Delivery,” Science Advances 7:eabd5835 (2021); and (28) Krishnan et al., “A Wireless, Battery-Free Device Enables Oxygen Generation and Immune Protection of Therapeutic Xenotransplants in vivo,” Proc. Nat’l Acad. Sci. USA 120:e2311707120 (2023). The summary in Table 3 is restricted to systems utilizing primary islets or SC-β cells that demonstrate diabetes reversal for a duration exceeding two weeks in immunocompetent animal models

[0158] Recent advancements in cell encapsulation technology have increasingly focused on the development of encapsulation systems capable of providing oxygenation to preserve viability and function of encapsulated cells (Pedraza et al., “Preventing Hypoxia-Induced Cell Death in Beta Cells and Islets via Hydrolytically Activated, Oxygen-Generating Biomaterials,” Proc. Natl. Acad. Sci. U.S.A.109(11):4245-50 (2012); Coronel et al., “Oxygen Generating Biomaterial Improves the Function and Efficacy of Beta Cells within a Macroencapsulation Device,” Biomaterials 210:1-11 (2019); Wang et al., “An Inverse-Breathing Encapsulation System for Cell Delivery,” Science Advances 7(20):eabd5835 (2021); Lee et al., “Electrocatalytic On-Site Oxygenation for Transplanted Cell-Based-Therapies,” Nature Communications 14(1):7019 320349050v3(2023); Krishnan et al., “A Wireless, Battery-Free Device Enables Oxygen Generation and Immune Protection of Therapeutic Xenotransplants in Vivo,” Proceedings of the National Academy of Sciences 120(40):e2311707120 (2023), which are hereby incorporated by reference in their entirety). For examples, metal peroxides (Pedraza et al., “Preventing Hypoxia-Induced Cell Death in Beta Cells and Islets via Hydrolytically Activated, Oxygen-Generating Biomaterials,” Proc. Natl. Acad. Sci. U.S.A.109(11):4245-50 (2012); Coronel et al., “Oxygen Generating Biomaterial Improves the Function and Efficacy of Beta Cells within a Macroencapsulation Device,” Biomaterials 210:1-11 (2019); Liang et al., “Engineering a Macroporous Oxygen-Generating Scaffold for Enhancing Islet Cell Transplantation within an Extrahepatic Site,” Acta Biomater 130:268-280 (2021); Harrison et al., “Oxygen Producing Biomaterials for Tissue Regeneration,” Biomaterials 28(31):4628-34 (2007); Wang et al., “An Inverse-Breathing Encapsulation System for Cell Delivery,” Science Advances 7(20):eabd5835 (2021), which are hereby incorporated by reference in their entirety) have been employed to generate oxygen to enhance cell survival in several encapsulation platforms. However, the inherently short-term oxygen supply in these methods poses a significant barrier to clinical translation, which demands long-term oxygenation. Thus, the need for a method to provide a continuous and sustainable oxygen source remains critical. A recent proof-of-concept study by Krishnan et al. successfully demonstrated the feasibility of using electrolysis-based oxygenation to improve the survival of encapsulated cells in subcutaneous space (Krishnan et al., “A Wireless, Battery-Free Device Enables Oxygen Generation and Immune Protection of Therapeutic Xenotransplants in Vivo,” Proceedings of the National Academy of Sciences 120(40):e2311707120 (2023), which is hereby incorporated by reference in its entirety). Their device, powered and regulated by resonant inductive coupling, effectively supported a moderate density of pancreatic islets (1000 IEQ / cm2) to correct diabetes in diabetic mice for a month. However, the practicality of resonant inductive coupling in clinical settings for generating an adequate level of oxygen to supply a metabolically relevant dose of cells requires further development and investigation.

[0159] A bioelectronics-assisted encapsulation (BEAM) system incorporating a built-in battery and electronic controller, capable of generating oxygen in a continuously regulated and precise manner was developed. The BEAM system is designed to overcome the limitations of oxygen supply duration associated with chemical-based methods by providing a theoretically unlimited oxygen supply through the electrolysis of tissue moisture, powered by a rechargeable energy source. The oxygen generator, with compact dimensions (13 mm diameter x 3.1 mm thickness) at its full scale, was proven to provide a sustained and adequate oxygen supply to 320349050v3support the upper limits of clinically relevant islet doses (600,000 IEQ-740,000 IEQ) for 11 months to 2.5 years in vitro and three months in vivo. The battery could be designed to be recharged weekly in a wireless fashion via transcutaneous energy transfer (TET), an established technology already in use for FDA cleared rechargeable neurostimulators, thereby reducing the need for patient intervention and compliance requirements. Furthermore, the oxygen generation rate is programmable and precisely controlled, allowing for adaptation to various cell doses and types with differing oxygen consumption rates.

[0160] Membrane electrode assembly (MEA) technology was used for the electrolysis of water to facilitate oxygen generation in this system. The MEA comprises a proton exchange membrane that, upon hydration, functions as the electrolyte, enabling proton transfer in the balanced electrochemical reaction. Hydrogen and oxygen evolve on opposite sides of the membrane and are transported through separate gas ports via convection, with less than 1% diffusive crossover, thereby minimizing the risk of spark ignition within the gas phases. The exterior of the iEOG is electrically insulated from the internal components, ensuring the absence of current or voltage hazards upon direct contact. Furthermore, the electrocatalyst-containing electrodes remain isolated from body fluids, as they receive water exclusively in vapor form through a vapor transport membrane, thereby mitigating any potential safety risks.

[0161] The linear outline of the BEAM system facilitates minimally invasive implantation and retrieval through a small skin incision. The cell encapsulation compartment is designed as an edge-free, flexible, round-ended cylindrical pouch, which minimizes the risk of enhanced fibrotic reactions typically caused by rigid, sharp edges. A straightforward sequential method for cell loading and system sealing was also developed, streamlining the preparation process, particularly under aseptic conditions. The concentric design of the cell encapsulation pouch, wherein cells are loaded into the thin annular space between the immune-protective membrane and the oxygen source (oxygen-permeable tubing), promotes optimal mass transport and ensures rapid, uniform oxygen diffusion. In addition, this structural configuration is scalable both radially and longitudinally, allowing for loading high islet doses while maintaining proximity between the encapsulated cells and the implant surface as well as the oxygen source.

[0162] The initial results demonstrated that oxygenation provided by the BEAM system successfully preserved the viability and insulin secretion of INS-1 aggregates and human pancreatic islets under a severe hypoxic in vitro condition (1% oxygen or 7.65 mmHg). Rats were selected as the animal model due to the greater thickness of the subcutaneous fat layer, which is more predictive as compared to mouse models when resembling the characteristic of human subcutaneous space (Grattoni et al., “Harnessing Cellular Therapeutics for Type 1 320349050v3Diabetes Mellitus: Progress, Challenges, and the Road Ahead,” Nature Reviews Endocrinology 21:14–30 (2025), which is hereby incorporated by reference in its entirety). Additionally, larger animals require higher dose of islets to correct diabetes, making it feasible to load them at a high density. In vivo studies showed that allogeneic islets encapsulated within the BEAM system successfully reversed diabetes and maintained normoglycemia for over one month and up to 88 days, when experiments were prematurely stopped for logistic reasons (i.e. rats compromising the oxygen transport tubing and electrical wire). These results were achieved at a high cell loading density (60,000 IEQ / mL, equivalent to approximately 4,200 IEQ / cm²). Meanwhile, islets encapsulated in pouches without oxygenation exhibited significant cell death and predominant loss of function. These findings provide proof-of-concept evidence for the feasibility of using oxygenation to maintain islet viability and function at high cell loading densities within poorly vascularized subcutaneous spaces. The loading density of 60,000 IEQ / mL was selected to achieve a reasonable dose (6000 IEQ) of islets to cure diabetes in rats (~11,000-15,000 IEQ / kg) using a practically small volume of cell suspension (100 μL). While the packing of 60,000 IEQ / mL or 4200 IEQ / cm2is among the highest densities reported in literature (Table 3), even higher density packing may be possible in clinical application of the BEAM system, especially considering that rat pancreatic islets have a 2-3-fold higher oxygen consumption rate compared to human islets. It was estimated that a pouch with an outer diameter of 1 cm and a length of 15 cm would be adequate for encapsulation of approximately 400,000 IEQ at a concentration of 120,000 IEQ / mL (Table 4, FIG.34). Alternatively, using seven pouches with an outer diameter of 0.75 cm and a length of 7 cm arranged in parallel could achieve the delivery of 420,000 human IEQ at a density of 60,000 IEQ / mL. The final dimensions of the system including the iEOG and connectors would approximate the size of a credit card (FIG.31). As smaller pancreatic islets (100-150 μm) have been shown to be superior to random-sized pancreatic islets (50-400 μm) in terms of their function (Ernst et al., “A Predictive Computational Platform for Optimizing the Design of Bioartificial Pancreas Devices,” Nature Communications 13(1):6031 (2022); Lehmann et al., “Superiority of Small Islets in Human Islet Transplantation,” Diabetes 56(3):594-603 (2007); Farhat et al., “Small Human Islets Comprised of More β-cells with Higher Insulin Content Than Large Islets,” Islets 5(2):87-94 (2013), which are hereby incorporated by reference in their entirety), the use of smaller sizes may be preferable in preparation of SC-β cell aggregate. It was estimated that a system of about half the size of a credit card could be enough to house 500,000 cell aggregates with diameters of 100 - 125 μm. (FIGs.32A-32B and 33). 320349050v3Table 4. The Capacity Estimation and the Corresponding Cell Doses that can be Loaded Into a Cell Encapsulation Pouch of the BEAM Device

[0163] Although the findings from the current design of the BEAM system are promising, several technical challenges must be overcome to realize its potential for clinical translation. Due to the relatively large size of the current electrical controller designed for bench testing, the bioelectronic components could not be fully implanted in rats. Thus, an extracorporeal setup was used, wherein the oxygen generator and electronic components were housed in an external pocket worn by the animal. This setup allowed for facile monitoring of iEOG function as well as tracking power and water consumption, which are valuable for the further development of the BEAM system. A more compact system has been designed, but not yet implemented, for use in humans. For example, in a clinical context, the entire system—including the cell encapsulation pouch, iEOG, battery, and electrical controller—would need to be fully implantable, with the battery recharged weekly via TET, as is currently used in implantable neurostimulators, left ventricular assist devices and others (Pya et al., “First Human Use of a Wireless Coplanar Energy Transfer Coupled with a Continuous-Flow Left Ventricular Assist Device,” The Journal of Heart and Lung Transplantation 38(4):339-343 (2019); Slaughter et al., “Transcutaneous Energy Transmission for Mechanical Circulatory Support Systems: History, Current Status, and Future Prospects,” Journal of Cardiac Surgery 25(4):484-489 (2010); Dissanayake et al., “Transcutaneous Energy Transfer System for Powering Implantable Biomedical Devices,” in 13th International Conference on Biomedical Engineering Berlin, Heidelberg: Springer Berlin Heidelberg (2009); Dual et al., “The Future of Durable Mechanical Circulatory Support: Emerging Technological Innovations and Considerations to Enable Evolution of the Field,” Journal of Cardiac Failure 30(4):596-609 (2024); Campi et al., “Wireless Power Transmission for Left Ventricular Assist Devices: Advancements, Challenges, and Future Directions,” in 2024 320349050v3IEEE Wireless Power Technology Conference and Expo (WPTCE) (2024), which are hereby incorporated by reference in their entirety). It was acknowledged that using an extracorporeal system, which relies on transcutaneous tubing and an external jacket to secure the electronic components is troublesome, particularly for long-term studies in flexible animals like rats. In fact, experiments were prematurely terminated in 6 out of 9 rats due to damaged oxygen transportation tubing or rats escaping from jackets (Table 2). Furthermore, although the jackets were initially designed to fit comfortably, the rats could outgrow them within a couple of months due to continuous weight gain as they returned to normoglycemia. The use of larger animals, such as pigs, in conjunction with a fully implantable system, is needed to further assess the long- term efficacy of the BEAM system. Second, it was envisioned that in a fully implantable system, it is necessary to develop a strategy to safely dissipate the hydrogen co-product generated during electrolysis from the body. Although hydrogen has been demonstrated to be safe in other contexts (Ohsawa et al., “Hydrogen Acts as a Therapeutic Antioxidant by Selectively Reducing Cytotoxic Oxygen Radicals,” Nature Medicine 13(6):688-694 (2007); Ge et al., “Molecular Hydrogen: a Preventive and Therapeutic Medical Gas for Various Diseases,” Oncotarget 8(60):102653-102673 (2017); Wu et al., “Hydrogen Gas from Inflammation Treatment to Cancer Therapy,” ACS Nano 13(8):8505-8511 (2019); Saengsin et al., “Hydrogen Therapy as a Potential Therapeutic Intervention in Heart Disease: From the Past Evidence to Future Application,” Cellular and Molecular Life Sciences 80(6):174 (2023); Tamura et al., “Efficacy of Inhaled Hydrogen on Neurological Outcome Following Brain Ischaemia During Post-Cardiac Arrest Care (HYBRID II): a Multi-Centre, Randomised, Double-Blind, Placebo-Controlled Trial,” eClinicalMedicine 58:101907 (2023), which are hereby incorporated by reference in their entirety), such as in scavenging reactive oxygen species, further investigation is required to assess the long-term impact of hydrogen produced by the BEAM system. A study by Zhu et al. demonstrated that hydrogen can be rapidly cleared from rat tissues within 25 minutes, even at saturated levels (Zhu et al., “Accurate in vivo Real-Time Determination of the Hydrogen Concentration in Different Tissues of Mice After Hydrogen Inhalation,” Heliyon 8(10):e10778 (2022), which is hereby incorporated by reference in its entirety), supporting the feasibility of safely dissipating hydrogen produced by the iEOG. The simulations indicate that hydrogen generation occurs at a rate of 0.13 scch (5.8 µmol / h) at 280 µA and 4.95 scch (221 µmol / h) at 11 mA. Given the reported clearance rate, these levels are expected to be safely dissipated through an appropriately designed tissue interface facilitating hydrogen diffusion. One potential strategy to enhance hydrogen clearance involves incorporating a porous membrane at the hydrogen 320349050v3outlet, thereby providing sufficient surface area for controlled diffusion of hydrogen in its soluble phase into the surrounding tissue and circulation.

[0164] Third, although the accumulation of excessive oxygen around the cell encapsulation pouch did not appear to adversely affect the encapsulated cells, it resulted in a separation of the system from the surrounding tissue in some cases, thereby hindering mass transport. Interestingly, the oxygen production in this study was measured at approximately 2.43 pmol / IEQ / min, a level comparable to or even lower than reported values for rat pancreatic islets in common culture conditions (Table 1). It is noteworthy that over 90% of the encapsulated islets remained viable and functional, exhibiting responses to glucose changes and maintaining the expression of insulin and glucagon, at the time the excessive gas phase was observed. Given that the oxygen consumption rates of beta cells are variable and highly dependent on environmental factors such as glucose and oxygen levels, it is plausible that the total oxygen produced at a constant rate of 2.43 pmol / IEQ / min would exceed the total oxygen demands of these islets under dynamic conditions. Furthermore, as the implant becomes gradually integrated into the subcutaneous space with newly formed vasculature, the oxygen demand may be less compared to the early post-implantation period. In fact, different levels of FBR and vascularization were observed around the cell encapsulation pouches; thus, the oxygen demands of islets in these implants may vary (FIG.35). This challenge could potentially be mitigated by incorporating a built-in sensor (e.g. total pressure) within the cell encapsulation pouch, thereby creating a closed- loop system capable of delivering “on-demand” oxygen to the encapsulated cells, as described previously in our patents (U.S. Patent Application Publication No.2018 / 0135948 to Stone et al.; U.S. Patent No.12,090,300 to Stone S., which are hereby incorporated by reference in their entirety).

[0165] The preceding examples underscores the pivotal importance of adequate oxygenation in encapsulation systems to deliver therapeutic cells. Enhanced oxygen levels efficiently sustained the viability and functionality of encapsulated insulin-secreting cells in both in vitro and in vivo contexts. Notably, continuous oxygen supply allows for the packing of these cells at higher densities without compromising their viability and function. This advancement holds promise for potentially minimizing the required size of encapsulation systems to levels suitable for clinical application, harnessing cell therapy for type 1 diabetes without the need for immunosuppression.

[0166] Although preferred embodiments have been depicted and described in detail herein, it will be apparent to those skilled in the relevant art that various modifications, additions, substitutions, and the like can be made without departing from the spirit of the invention and 320349050v3these are therefore considered to be within the scope of the invention as defined in the claims which follow. 320349050v3

Claims

WHAT IS CLAIMED:

1. An implantable therapeutic delivery system comprising: an electrochemical oxygen generator; a cell compartment comprising a semi-permeable nanofibrous membrane that is impermeable to cellular migration, an internal hydrogel matrix and one or more cells embedded in the hydrogel matrix; a conduit in fluid communication between the electrochemical oxygen generator and the interior of the cell compartment, wherein the conduit delivers oxygen generated by the electrochemical oxygen generator to the one or more cells embedded in the hydrogel matrix.

2. The implantable therapeutic delivery system of claim 1, wherein the conduit comprises a first portion that is impermeable to oxygen and a second portion that is permeable to oxygen.

3. The implantable therapeutic delivery system of claim 2, wherein the first portion is primarily located externally of the cell compartment and the second portion is entirely located internally to the cell compartment.

4. The implantable therapeutic delivery system of claim 3, wherein the second portion of the conduit is located centrally within the cell compartment and extends the full length of the cell compartment.

5. The implantable therapeutic delivery system of claim 3, wherein a terminal end of the second portion of the conduit is sealed.

6. The implantable therapeutic delivery system of claim 2 further comprising a hermetical seal formed at the junction of the cell compartment membrane, and the first and second portions of the conduit.

7. The implantable therapeutic delivery system of claim 1, wherein the semi- permeable nanofibrous membrane is formed by electrospinning.

8. The implantable therapeutic delivery system of claim 1, wherein the semi- permeable nanofibrous membrane is formed of nanofibers comprising one or more medical- grade elastomers. 320349050v39. The implantable therapeutic delivery system of claim 8, wherein the one or more medical-grade elastomers are selected from the group of polyethylene glycol, polyamide, polyether block amide, polycarbonate urethane, thermoplastic silicon-polycarbonate-urethane, polyether urethane, and combinations thereof.

10. The implantable therapeutic delivery system of any one of claims 1 to 9, wherein the semi-permeable nanofibrous membrane is seamless.

11. The implantable therapeutic delivery system of any one of claims 1 to 10, wherein the semi-permeable nanofibrous membrane further comprises one or more nitinol mesh layers.

12. The implantable therapeutic delivery system of any one of claims 1 to 11, wherein the semi-permeable nanofibrous membrane is either (i) a hydrogel-reinforced nanofibrous membrane, (ii) a nitinol mesh-reinforced nanofibrous membrane, or (iii) a hydrogel and nitinol mesh-reinforced nanofibrous membrane.

13. The implantable therapeutic delivery system of any one of claims 1 to 12, wherein the hydrogel material comprises a natural polymeric material, a synthetic polymeric material, or a combination thereof.

14. The implantable therapeutic delivery system of claim 1 or 13, wherein the hydrogel material comprises a natural polymeric material selected from the group consisting of collagen, hyaluronate, fibrin, alginate, agarose, chitosan, bacterial cellulose, elastin, keratin, derivatives thereof, and combinations thereof.

15. The implantable therapeutic delivery system of claim 14, wherein the hydrogel material comprises a pure alginate, a modified alginate, or a mixture of pure and modified alginate.

16. The implantable therapeutic delivery system of claim 15, wherein the modified alginate is a zwitterionically modified alginate.

17. The implantable therapeutic delivery system of claim 1 or 13, wherein the hydrogel material comprises a synthetic polymeric material selected from polyethylene glycol (PEG), poly(acrylic acid), poly(ethylene oxide), poly(vinyl alcohol), polyphosphazene, poly(hydroxyethyl methacrylate), triazole-zwitterion hydrogels (TR-qCB, TR-CB, TR-SB), 320349050v3poly(sulfobetaine methacrylate), carboxybetaine methacrylate, poly[2-methacryloyloxyethyl phosphorylcholine, N-hydroxyethyl acrylamide, a copolymer thereof, a derivatives thereof, and a combination thereof.

18. The implantable therapeutic delivery system according to claim 12, wherein the nitinol mesh-reinforced nanofibrous material comprises one or more nanofiber layers, such as first and second nanofibrous layers with the nitinol mesh sandwiched between the first and second nanofibrous layers, optionally wherein fibers of the first and second nanofibrous layers are bonded to one another via pores in the nitinol mesh.

19. The implantable therapeutic delivery system of one of claims 12 or 18, wherein the nitinol mesh-reinforced nanofibrous material has a thickness of about 10 to about 100 μm, about 10 to about 50 μm, or about 50 to about 100 μm, such as about 10 to about 20 μm, about 20 to about 30 μm, about 30 to about 40 μm, about 40 to about 50 μm, about 50 to about 60 μm, about 60 to about 70 μm, about 70 to about 80 μm, about 80 to about 90 μm, about 90 to about 100 μm.

20. The implantable therapeutic delivery system of any one of claims 1 to 19, wherein the hydrogel matrix is a natural polymeric material, a synthetic polymeric material, or a combination thereof, as defined in one of claims 14 to 17.

21. The implantable therapeutic delivery system of any one of claims 1 to 20, wherein the cells positioned within the hydrogel matrix secrete at least one therapeutic agent.

22. The implantable therapeutic delivery system of any one of claims 1 to 21, further comprising a second therapeutic agent positioned within the hydrogel material.

23. The implantable therapeutic delivery system of any one of claims 1 to 22, wherein the preparation of cells comprises a preparation of single cells or a preparation of cell aggregates.

24. The implantable therapeutic delivery system of any one of claims 1 to 23, wherein the preparation of cells comprises a preparation of primary cells, a preparation of stem-cell derived cells, or a preparation of immortalized cells.

25. The implantable therapeutic delivery system of any one of claims 1 to 24, wherein the preparation of cells comprise a preparation of mammalian cells. 320349050v326. The implantable therapeutic delivery system of claim 25, wherein the preparation of cells comprise a preparation of mammalian cells selected from the group consisting of primate cells, rodent cells, canine cells, feline cells, equine cells, bovine cells, and porcine cells.

27. The implantable therapeutic delivery system of claim 26, wherein the preparation of cells comprise human cells.

28. The implantable therapeutic delivery system of claim 24, wherein the preparation of cells comprise a preparation of stem cell derived cells.

29. The implantable therapeutic delivery system of claim 28, wherein the stem cell derived cells are derived from pluripotent, multipotent, oligopotent, or unipotent stem cells.

30. The implantable therapeutic delivery system of claim 28, wherein the stem cell derived cells are derived from embryonic stem cells, epiblast cells, primitive ectoderm cells, primordial germ cells, or induced pluripotent stem cells.

31. The implantable therapeutic delivery system of any one of claims 1 to 27, wherein the preparation of cells comprise cells selected from the group consisting of smooth muscle cells, cardiac myocytes, platelets, epithelial cells, endothelial cells, urothelial cells, fibroblasts, embryonic fibroblasts, myoblasts, chondrocytes, chondroblasts, osteoblasts, osteoclasts, keratinocytes, hepatocytes, bile duct cells, islets or islet cells, thyroid, parathyroid, adrenal, hypothalamic, pituitary, ovarian, testicular, salivary gland cells, adipocytes, embryonic stem cells, mesenchymal stem cells, neural cells, endothelial progenitor cells, hematopoietic cells, precursor cells, mesenchymal stromal cells, Baby Hamster Kidney (BHK) cells, Chinese Hamster Ovary cells, Human Amniotic Epithelial (HAE) cells, choroid plexus cells, chromaffin cells, adrenal chromaffin cells, pheochomocytoma cell line PC12, human retinal pigment epithelium cells, recombinant human retinal pigment epithelium cells, NGF-secreting Baby Hamster Kidney (BHK) cells, human bone marrow-derived stem cells transfected with GLP-1, BDNF-producing fibroblasts, NGF-producing cells, CNTF-producing cells, BDNF-secreting Schwann cells, IL-2-secreting myoblasts, endostatin-secreting cells, and cytochrome P450 enzyme overexpressed feline kidney epithelial cells, myogenic cells, embryonic stem cell- derived neural progenitor cells, irradiated tumor cells, proximal tubule cells, neural precursor cells, astrocytes, genetically engineered cells. 320349050v332. The implantable therapeutic delivery system of claim 31, wherein the preparation of cells comprises a preparation of islets or a preparation of islet cells that release insulin and glucagon.

33. The implantable therapeutic delivery system of claim 32, wherein the preparation of islets is a preparation of primate islets, rodent islets, canine islets, feline islets, equine islets, bovine islets, or porcine islets.

34. The implantable therapeutic delivery system of claim 32 or 33, wherein the preparation of islets comprises a density between 1x103to 2x106islet equivalents (IEQs) / mL.

35. The implantable therapeutic delivery system of any one of claims 21 or 22, wherein the therapeutic agent and / or the second therapeutic agent comprises one or more biologically active agents selected from the group consisting of a protein, peptide, antibody or antibody fragment thereof, antibody mimetic, a nucleic acid, a small molecule, a hormone, a growth factor, an angiogenic factor, a cytokine, an anti-inflammatory agent, an anti-fibrotic agent, and combinations thereof.

36. The implantable therapeutic delivery system of 22, wherein the second therapeutic agent comprises an anti-fibrotic agent.

37. The implantable therapeutic delivery system of claim 36, wherein the anti-fibrotic agent is in crystalline form.

38. The implantable therapeutic delivery system of claim 36, wherein the anti-fibrotic agent is nintedanib, GW2580, or pirfenidone.

39. The implantable therapeutic delivery device of any one of claims 1 to 36, wherein the conduit comprises silicone.

40. A method of delivering a therapeutic agent to a subject in need thereof, said method comprising: 320349050v3implanting the implantable therapeutic delivery system according to any one of claims 1- 39 into the subject.

41. A method of treating diabetes in a subject, said method comprising: implanting the implantable therapeutic delivery system according to any one of claims 32 to 34 into the subject having diabetes.

42. The method of claim 41, wherein the hydrogel matrix comprises a preparation of cells that release insulin, glucagon, or a combination thereof for the treatment of diabetes in the subject.

43. The method of claim 41 or 42, wherein the preparation of cells is a preparation of islets.

44. The method of claim 43, where the preparation of islets is a preparation of primate islets, rodent islets, canine islets, feline islets, equine islets, bovine islets, or porcine islets.

45. The method of claim 41 or 42, wherein the preparation of cells is an islet-like cluster derived from a preparation of stem cells.

46. The method of claim 43, wherein the preparation of stem cells is selected from the group consisting of embryonic stem cells, epiblast cells, primitive ectoderm cells, primordial germ cells, and induced pluripotent stem cells.

47. A method of treating a bleeding disorder in a subject, said method comprising: implanting the implantable therapeutic delivery system of any one of claims 1 to 31, 35 or 39 into the subject having a bleeding disorder.

48. The method of claim 47, wherein the bleeding disorder is selected from the group consisting of hemophilia A, hemophilia B, von Willebrand disease, Factor I deficiency, Factor II deficiency, Factor V deficiency, Factor VII deficiency, Factor X deficiency, Factor XI deficiency, Factor XII deficiency, and Factor XIII deficiency. 320349050v349. The method of claim 47 or 48, wherein the hydrogel matrix comprises a preparation of cells that release one or more blood clotting factors selected from the group consisting of Factor I, Factor II, Factor V, Factor VII, Factor VIII, Factor IX, Factor X, Factor XI, Factor XII, and Factor XIII for treatment of the bleeding disorder.

50. The method of claim 47, wherein the preparation of cells comprises recombinant myoblasts, mesenchymal stromal cells, induced pluripotent stem cell derived endothelial cells, or a combination thereof.

51. A method of treating a lysosomal storage disease in a subject, said method comprising: implanting the implantable therapeutic delivery system of any one of claims 1 to 31, 35 or 39 into the subject having the lysosomal storage disease.

52. The method of claim 51, wherein the hydrogel matrix comprises a preparation of cells that release an enzyme selected from the group consisting of α-L-iduronidase, Iduronate-2- sulfatase, α-glucuronidase, Arylsulfatase A, alpha-Galactosidase A, and combinations thereof, for treating the lysosomal storage disease in the subject.

53. The method of claim 52, wherein the preparation of cells comprises hematopoietic stem cells, fibroblasts, myoblasts, Baby Hamster Kidney (BHK) cells, Chinese Hamster Ovary cells, Human Amniotic Epithelial (HAE) cells, or combinations thereof.

54. A method of treating a neurological disorder in a subject, said method comprising: implanting the implantable therapeutic delivery system of any one of claims 1 to 31, 35 or 39 into the subject having the neurological disorder.

55. The method of claim 54, wherein the neurological disorder is selected from the group consisting of Parkinson’s disorder, Alzheimer’s disease, epilepsy, Huntington’s disease, Amyotrophic lateral sclerosis, chronic pain, visual loss, hearing loss, peripheral nerve injury, and spinal cord injury. 320349050v356. The method of claim 54 or 55, wherein the hydrogel matrix comprises a preparation of cells that release a molecule selected from cerebrospinal fluid, extracellular fluid, levodopa, nerve growth factor (NGF), ciliary neurotrophic factor (CNTF), BLP-1, brain-derived neurotrophic factor (BDNF), vascular endothelial growth factor (VEGF), enkephalin, adrenaline, catecholamine, and combinations thereof, for treating the neurological disorder.

57. The method of claim 56, wherein the preparation of cells comprises choroid plexus cells, chromaffin cells, pheochomocytoma cell line PC12, human retinal pigment epithelial cells, NGF-secreting Baby Hamster Kidney (BHK) cells, myoblasts, human bone marrow-derived stem cells transfected with GLP-1, BDNF-producing fibroblasts, NGF- producing cells, CNTF-producing cells, adrenal chromaffin cells, BDNF-secreting Schwann cells, myogenic cells, embryonic stem cell-derived neural progenitor cells, or combinations thereof.

58. A method of treating a cancer in a subject, said method comprising: implanting the implantable therapeutic delivery system of any one of claims 1 to 31, 35 or 39 into the subject having cancer.

59. The method of claim 58, wherein the hydrogel matrix comprises a preparation of cells that release a molecule selected from IL-2, endostatin, cytochrome P450 enzyme, tumor antigens, a cytokine, and combinations thereof, for treating cancer in the subject.

60. The method of claim 59, wherein the preparation of cells comprises IL-2- secreting myoblasts, endostatin-secreting cells, Chinese Hamster Ovary cells, and cytochrome P450 enzyme overexpressed feline kidney epithelial cells, irradiated tumor cells, or combinations thereof.

61. A method of treating a chronic eye disease in a subject, said method comprising: implanting the implantable therapeutic delivery system of any one of claims 1 to 31, 35 or 39 into the subject having a chronic eye disease.

62. The method of claim 61, wherein the chronic eye disease is selected from the group consisting of age-related macular degeneration, diabetic retinopathy, retinitis pigmentosa, glaucoma, macular telangiectasia, and combinations thereof. 320349050v363. The method of claim 61 or 62, wherein the hydrogel matrix comprises a preparation of cells that release a molecule selected from ciliary neurotrophic factor, antagonists against vascular endothelial growth factor and platelet-derived growth factor, and combinations thereof, for treating the chronic eye disease.

64. The method of claim 63, wherein the preparation of cells comprises human retinal pigment epithelium cells, recombinant human retinal pigment epithelium cells, or a combination thereof.

65. A method of treating a kidney failure in a subject, said method comprising: implanting the implantable therapeutic delivery system of any one of claims 1 to 31, 35 or 39 into the subject having a kidney failure.

66. The method of claim 65, wherein the hydrogel matrix comprises a preparation of cells that release a therapeutic molecule suitable for treating the kidney failure.

67. The method of claim 65 or 66, wherein the hydrogel matrix comprises a preparation of renal proximal tubule cells, mesenchymal stem cells, and a combination thereof.

68. A method of treating a chronic pain in a subject, said method comprising: implanting the implantable therapeutic delivery system of any one of claims 1 to 31, 35 or 39 into the subject having a chronic pain.

69. The method of claim 68, wherein the chronic pain is chronic pain caused by degenerative joint, peripheral neuropathy, or cancer.

70. The method of claim 68 or 69, wherein the hydrogel matrix comprises a preparation of cells that release a molecule selected from the group consisting of catecholamine, opioid peptides, enkephalins, and combinations thereof.

71. The method of claim 70, wherein the preparation of cell comprises chromaffin cells, neural precursor cells, mesenchymal stem cells, astrocytes, and genetically engineered cells, or a combination thereof. 320349050v372. The method according to any one of claims 40 to 71, wherein said implanting is carried out via a laparoscopic procedure.

73. The method according to any one of claims 40 to 71, wherein said implantable therapeutic delivery system is implanted intraperitoneally, percutaneously, or subcutaneously.

74. The method according to any one of claims 40 to 71, wherein said method further comprises retrieving the implantable therapeutic delivery system from the subject.

75. The method according to claim 74, wherein said method further comprises implanting a replacement implantable therapeutic delivery system after said retrieving.

76. The method according to any one of claims 40 to 73 further comprising replacing a battery of the electrochemical oxygen generator without retrieving the implantable therapeutic delivery system.

77. A method of forming a seamless nanofibrous membrane comprising: forming an enlarged diameter support about a mandrel, the support being formed of a dissolvable material; and electrospinning a nanofibrous membrane about the enlarged diameter support and a portion of the mandrel.

78. The method of claim 75 further comprising: dissolving the enlarged diameter support; and removing the seamless nanofibrous membrane from the mandrel.

79. A method of making a cell encapsulation system suitable for implant, the method comprising: providing a semi-permeable nanofibrous membrane structure; inserting a permeable conduit into the semi-permeable nanofibrous membrane, the permeable conduit having an unsealed end; introducing cells and hydrogel precursor via the permeable conduit, whereby the cells and hydrogel precursor are forced into the space between the permeable conduit and the 320349050v3nanofibrous membrane, followed by crosslinking of the hydrogel precursor to form the hydrogel matrix; joining together the unsealed end of the permeable conduit to a distal end of an impermeable conduit coupled to an electrochemical oxygen generator; and sealing the semi-permeable nanofibrous membrane about the distal end of the impermeable conduit.

80. The method according to claim 79, wherein the semi-permeable nanofibrous membrane structure has a single opening, and the permeable conduit is inserted through the single opening.

81. The method according to claim 79 or 80, wherein said sealing comprises adhesively sealing the semi-permeable nanofibrous membrane about the distal end of the impermeable conduit.

82. The method according to claim 79 or 80, wherein said sealing comprises installing a hermetic coupling between the semi-permeable nanofibrous membrane and the distal end of the impermeable conduit.

83. A device for forming an implantable therapeutic delivery system comprising: an electrochemical oxygen generator; a compartment comprising a semi-permeable nanofibrous membrane that is impermeable to cellular migration; a conduit in fluid communication between the electrochemical oxygen generator and the interior of the compartment, wherein the conduit delivers oxygen generated by the electrochemical oxygen generator to the interior of the compartment.

84. A kit comprising: the device of claim 83; one or more precursors for forming a hydrogel matrix; and instructions for loading a composition comprising cells and the one or more precursors into the compartment of the device, and for forming the hydrogel matrix in situ. 320349050v3

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