Tunable degradation of hydrogel microparticles

Abstract

Non-alginate hydrogel microparticles for the localized delivery and sustained release of therapeutic cells and / or tissue (including homogeneous or heterogeneous cell masses) at an implantation site. The microparticles comprise a three-dimensional matrix of a covalently crosslinked non-alginate polymeric compound and a therapeutically effective amount of cells and / or tissue encapsulated therein, the cells having at least about 50% viability, and the microparticles having a size greater than about 30 μm. Compositions containing such microparticles and methods of using such microparticles for therapy are also described.

Description

[Technical field]

[0001] CROSS-REFERENCE TO RELATED APPLICATIONS This application claims the benefit of priority to U.S. Provisional Patent Application No. 62 / 858,578, filed June 7, 2019, entitled Tunable Degradation of Hydrogel Microparticles, which is incorporated by reference in its entirety. Technical Field

[0002] The present invention relates to hydrogel-based microparticles for encapsulating biologics, as well as methods for adjusting the physical and chemical characteristics of the hydrogel matrix, such as to increase or decrease the degradation profile of each microparticle. [Background technology]

[0003] 2. Description of Related Art Cell microencapsulation is a rapidly expanding field that has the potential to show broad utility in the treatment of diseases such as diabetes, cancer, heart disease, and immune deficiencies, as well as in tissue engineering research and regenerative medicine. Traditional alginate microspheres still suffer from poor biocompatibility, and microencapsulation using more advanced hydrogels suffers from slow gelation rates. There is a need for non-cytotoxic, non-emulsion-based hydrogel microparticles that can be produced from a wide variety of hydrogel materials. Summary of the Invention

[0004] Described herein are non-alginate hydrogel microparticles for localized delivery and sustained release of therapeutic cells and / or tissues (including homogeneous or heterogeneous cell masses) at an implantation site. The microparticles include a three-dimensional matrix of covalently crosslinked non-alginate polymeric compounds and a therapeutically effective amount of cells and / or tissues encapsulated therein, the cells having at least 50% viability, and the microparticles have a size of greater than about 30 μm. Exemplary polymeric compounds are selected from slow gelling polymer precursors selected from the group consisting of branched or unbranched hyaluronic acid, branched or unbranched functionalized hyaluronic acid, branched or unbranched functionalized polyethylene glycol, hyaluronan, fibrin, chitosan, collagen, polylactic acid, poly(L-lactic acid), polylactic-co-glycolic acid, polycaprolactone, polyvinyl alcohol, and combinations thereof. The matrix may further comprise a crosslinker crosslinked with the non-alginic polymeric compound, selected from the group consisting of dithiothreitol, branched or unbranched functionalized polyethylene glycol, dithiols, ethylene glycol bis-mercaptoacetate, and combinations thereof. Exemplary functionalized polyethylene glycols include, but are not limited to, polyethylene glycol dithiol, polyethylene glycol diacrylate, polyethylene glycol divinylsulfone, polyethylene glycol dimaleimide, and combinations thereof. The matrix may be a homogeneous matrix comprising one type of covalently crosslinked polymer backbone (with or without additional crosslinkers of different polymeric types). The matrix may also be a heterogeneous matrix comprising a mixture of two or more polymer precursors, such as a combination of high molecular weight (500 kDA or more, preferably 100 kDA or more) and low molecular weight (less than 100 kDA, preferably 50 kDa or less) polymer precursors, and / or a mixture of two or more crosslinkers.

[0005] Compositions containing a plurality of such microparticles are also described herein. Such compositions can also contain a plurality of "empty" microparticles dispersed along with the cell or tissue loaded microparticles. The compositions can contain a mixture of different microparticles (e.g., a mixture of degradable hydrogel microparticles and durable hydrogel microparticles), and / or a mixture of microparticles of different sizes (e.g., a plurality of large microparticles and a plurality of small microparticles), and / or a mixture of microparticles each containing a different type of cell and / or tissue (e.g., insulin producing cells in one set of microparticles and supporting stem cells in another set of microparticles). Two different compositions can be combined, or the two different types of particles can be co-administered without being mixed into a single composition.

[0006] Also contemplated herein are various methods for the localized delivery and sustained release of therapeutic cells and / or tissues at the site of implantation by administering the microparticles and / or compositions containing such microparticles (alone or, if necessary, in combination with other types of therapeutic compounds). Typically, the microparticles or compositions are injected or implanted directly at or near (adjacent) the treatment site.

[0007] Also described herein are various methods and techniques for tuning the degradation rate of the microparticles by varying various parameters of the microparticles, including changes in formulation and / or processing parameters, such as the mass fraction or molecular weight of the polymer precursor, the molecular weight of the crosslinker, the ratio of crosslinker to polymer precursor, crosslinker hydrolysis, crosslinking kinetics, crosslinking time, e.g., UV exposure time, and combinations thereof.

[0008] Additionally, embodiments described herein contemplate the use of the empty microparticles for joint lubrication and / or viscosupplementation therapy by implanting multiple empty microparticles into a joint. [Brief description of the drawings]

[0009] The patent or application file contains at least one color drawing. Copies of this patent or patent application publication containing color drawing(s) will be provided by the U.S. Patent and Trademark Office upon request and payment of the necessary fee.

[0010] [Figure 1] FIG. 1 illustrates an example of covalent cross-linking between hydrogel precursor compounds. [Diagram 2] FIG. 2 illustrates the process for forming the above microparticles, including (A) droplet formation, (B) incubation in an alginate bath, (C) formation and thickening of the alginate shell, and (D) removal of the alginate shell after crosslinking of the hydrogel core to generate hydrogel microparticles. [Diagram 3] FIG. 3 shows images of (A) the alginate shell surrounding the microparticles; and (B) the hydrogel microparticles after removal of the alginate shell. [Figure 4] FIG. 4 shows images of (A) durable spherical PEG-based microparticles loaded with islets, (B) ellipsoidal PEG-based microparticles loaded with islets, (C) flat-edged MeHA-based microparticles loaded with stem cells, and (D) ellipsoidal MeHA-based microparticles loaded with stem cells. [Diagram 5] FIG. 5 shows the results of cytotoxicity studies: (A) the effect of calcium concentration and exposure time on the viability of canine pancreatic islets, and (B) the effect of photoinitiator concentration and UV exposure time on cell viability. [Figure 6] FIG. 6 is a graph plotting the diameters of 100 representative microparticles composed of ThHA, MeHA, PEGDA, or AHA, showing their good monodispersity profiles. [Figure 7] FIG. 7 shows images of fluorescently labeled dextran diffused into PEGDA hydrogel microparticles, ThHA hydrogel microparticles, and MeHA hydrogel microparticles. [Figure 8A] FIG. 8A is a graph of the decomposition of various particles over time, based on the mass of particulate polymer remaining. [Figure 8B] FIG. 8B is a graph showing the change in Q value (swelling ratio) over time for various particles. [Figure 8C] FIG. 8C is a graph of the degradation of various particles over time, based on the loss of particle mass. [Figure 9] FIG. 9 is a graph of the average daily blood glucose levels for each group in the initial study cohort after transplantation in mice. [Figure 10] Figure 10 shows images obtained from recovery of microparticles from the intraperitoneal space (IP space) of mice, with viability shown by deep red dithizone staining for (A) PEDGA, (B) MeHA, and (C) PEDGA, and viability shown by deep red dithizone and green calcein co-staining for (D) PEDGA. [Figure 11] FIG. 11 is an image showing the absence of fibrous tissue in healthy adipocytes after implantation with PEGDA microparticles. [Figure 12] FIG. 12 is a graph of the mean non-fasting blood glucose in dogs from measurements taken each morning and afternoon. [Figure 13] Figure 13 shows (A) an image of HEK293 cells clustered within durable PEGDA microparticles, (B) an image of the PEGDA microparticles after 34 days showing little change from (A) in cell morphology or number, (C) an image of HEK293 cells dispersed within degradable MeHA microparticles, and (D) an image of cells proliferating from (C) within the MeHA microparticles after 34 days. [Figure 14] FIG. 14 shows images obtained from microplate wells containing harvested spent media, demonstrating (A) very few HEK293 cells diffused from PEGDA microparticles and (B) a high number of viable cells diffused from MeHA microparticles. [Figure 15] FIG. 15 is a graph summarizing the average data from the cell release accumulation study. [Figure 16]FIG. 16 shows (A) an image of MSCs encapsulated in PEGDA microparticles, (B) an image of MSCs encapsulated in MeHA microparticles, (C) an image of pancreatic islets encapsulated in PEGDA microparticles, (D) an image of pancreatic islets encapsulated in MeHA microparticles, and (E) an image of MSCs encapsulated in AHA microparticles. [Figure 17] FIG. 17 is a graph of cell viability when rat insulinoma cell lines are encapsulated in PEGDA and AHA microparticles. [Figure 18] FIG. 18 shows images of islets encapsulated in PEGDA or MeHA and stained with dithiozone to demonstrate functional islets producing insulin. [Figure 19] FIG. 19 shows (A) an image of PEDGA microparticles being injected into and around the ventral retina, (B) an image of the microparticles adhering to tissue in the first dog, and (C) an image of the microparticles adhering to tissue in the second dog, demonstrating the "stickiness" of the particles. [Figure 20] FIG. 20 shows (A) an image of blue-stained PEGDA microparticles visible through the muscle in a rat knee joint; (B) an image of a dissected knee joint showing blue-stained PEGDA microparticles in the knee joint capsule region; and (C) an image of a further dissected knee joint showing blue-stained PEGDA microparticles remaining in the knee joint where they were originally implanted. [Figure 21] FIG. 21 shows images of hematoxylin and eosin staining of explant tissue for the three microparticle populations. [Figure 22] FIG. 22 is a graph of the (lack of) foreign body response in the retina to each particle. [Diagram 23] FIG. 23 is a graph of the effect of the mass fraction of the core (backbone) polymer species used in the hydrogel precursor solution on the Q value. [Figure 24]Figure 24A is a graph showing a formulation where the Q value decreased with increasing crosslinker concentration, and Figure 24B is a graph showing a formulation where the Q value increased with increasing crosslinker concentration. [Diagram 25] Figure 25A is a graph showing a formulation that results in an increased Q value with increasing pH, and Figure 25B is a graph showing a formulation that results in a decreased Q value with increasing pH. [Figure 26] Figure 26A is a graph showing a formulation where the Q value increased with increasing crosslinker molecular weight, and Figure 26B is a graph showing a formulation where the Q value did not change with increasing crosslinker molecular weight. [Figure 27] Figure 27A is a graph of the effect of changing the molecular weight of the core polymer species on the Q value for a constant number of reactive sites / gram, and Figure 27B is a graph of the effect of changing the molecular weight of the core polymer species on the Q value for different numbers of reactive sites / gram. [Figure 28] Figure 28A is a graph of the effect of UV crosslinking time on the Q value (swelling ratio) and diameter of the particles for various microparticle formulations. Figure 28B is a graph of the effect of UV crosslinking time on the Q value of MeHA microparticles. DETAILED DESCRIPTION OF THE PREFERRED EMBODIMENTS

[0011] The present invention relates generally to hydrogel microparticles for the local delivery and sustained release of therapeutic cells and tissues. The hydrogel microparticles protect the cells after implantation from initial degradation and washout (biodegradation) under physiological conditions and modulate the release kinetics of the cells (and their therapeutic proteins and molecules) to a local or systemic region over a period of hours to weeks or months. Hydrogel microparticles designed to degrade will eventually undergo in vivo degradation by normal physiological processes over a period of days or weeks, usually less than three months. This means that the entire hydrogel has been degraded and washed out by normal physiological conditions within this period to such an extent that substantially no hydrogel remains in the implanted area (it should be understood that trace amounts of gel or polymer may remain in the area, but usually not enough anymore to act as an encapsulant for the implanted cells). It will be understood that in some indications, especially when implanted in a joint, the microparticles will also undergo mechanical degradation due to movement and impact at the joint site.

[0012] The hydrogel microparticles include a hydrogel matrix in which a therapeutically effective amount of cells are encapsulated. The cells can be eukaryotic or prokaryotic. Examples of prokaryotic cells include bacteria and archaea. Eukaryotic cells include animal cells, plant cells, and fungi. The cells can be non-proliferative or proliferative. Proliferative cells will further proliferate (increase) from the initial cell number encapsulated in the hydrogel before and / or during implantation, or after release from the degrading hydrogel. Non-proliferative cells have limited proliferation capacity in vitro and generally do not increase from the initial cell number. However, the conditions within the hydrogel microparticles are favorable such that cell death in the encapsulated cells is minimal and the initial cell number of non-proliferative cells encapsulated in the hydrogel is not significantly reduced. In one or more embodiments, the initial viability of the cells after encapsulation in the microparticles remains at least 50% after encapsulation, preferably at least about 70%, more preferably at least about 80%, even more preferably at least about 90%, even more preferably at least about 95% viability, when compared to the viability of the starting cell population contained in the precursor solution. In other words, less than 50% of the cell population is lost during encapsulation, and preferably less than 30%, more preferably less than 20%, even more preferably less than 90%, even more preferably less than 5% of the cell population is lost during the encapsulation process. This is because the encapsulation process is gentle and cell culture medium and nutrients can diffuse into the microparticles during storage. In other words, in one or more embodiments, the microparticles have a low diffusion barrier.

[0013] In one or more embodiments, the cell population contained in the hydrogel precursor solution is about 1×10 6 cells / mL of the hydrogel precursor solution. 6 ~Approx. 5×10 8 The cells will be contained in a volume of about 4 x 10 cells per mL of hydrogel precursor solution. 6 ~ approx. 3 x 10 cells 8cells per mL of precursor solution, and in some embodiments, about 5×10 6 ~Approx. 1.5×10 8 In one or more embodiments, the hydrogel precursor solution contains approximately 15% by volume of the cells, cell mass, or tissue to be encapsulated per mL of the hydrogel precursor solution.

[0014] Examples of proliferating cells include stem cells and activated T cells, while examples of non-proliferating cells include pancreatic islets and differentiated neural cells. In either case, the cells (and their associated cell products) are slowly released from the hydrogel into the local implantation area. Exemplary cell products that may be secreted with the cells include signaling molecules, therapeutic proteins, vesicles, antibodies, viruses, exosomes, and the like. These protein-based components can be in native form or genetically modified. Similarly, the cellular components of the products can be native or genetically modified. The hydrogel microparticles allow the cells to be maintained in the local implantation area for longer periods of time compared to other carriers or non-encapsulated cells, improving the therapeutic effect of the implanted cells by providing a temporary, but sustainable, release of the cells and / or their signaling molecules, proteins, and the like, into the area. It will be appreciated that cell loading depends on the size of the cells and the size of each microparticle used. In one or more embodiments, up to 50% of the microparticle volume is comprised of encapsulated cells, cell clumps, and / or tissue, and preferably the cell, cluster, and / or tissue loading is in the range of about 10% to about 20% of the volume of the microparticle body. Typically, each 1 mm microparticle contains about 50 to about 1×10 5 Each 1 mm microparticle can encapsulate about 100 to about 1 × 10 cells. 5 The individual is encapsulated.

[0015] In one or more embodiments, the microparticles are formed from weaker hydrogels that will eventually degrade in vivo within about three months (90 days) or less from the date of implantation, referred to herein as "degradable" hydrogels (with the understanding that most hydrogels will eventually degrade). As used herein, "degradable" hydrogel microparticles are defined as "sustained-release" yet "instantly degradable" or "short-term" hydrogels that maintain a self-supporting microparticle body for at least 24 hours in PBS and up to three months when implanted in vivo. In addition, such degradable hydrogel microparticles degrade and deteriorate within about six months under in vitro storage conditions (PBS, 37°C). In other words, the hydrogel matrix has an "in vitro storage stability" of less than six months, meaning that when stored in PBS at 37°C, the hydrogel matrix begins to hydrolyze and disintegrate from a self-supporting body within six months of storage. In one or more embodiments, the degradable hydrogel microparticles are characterized by having a Q value greater than 150, and preferably greater than 200. However, as shown below, by modifying the crosslinking (e.g., using easily hydrolyzable crosslinkers), it is possible to obtain microparticles with lower Q values ​​(stronger initial matrices), which still degrade fairly rapidly and are therefore considered degradable hydrogel microparticles.

[0016] These degradable microparticles are in contrast to another embodiment, which is directed to durable hydrogel microparticles. Such durable hydrogel microparticles are also sustained release, but for longer term applications, such as not disintegrating or degrading under storage conditions in PBS at 37° C. for at least 6 months. In other words, the durable microparticles have an "in vitro storage stability" of more than 6 months. In one or more embodiments, the durable hydrogel particles are capable of long-term storage in PBS at room temperature (27° C.) for more than 1 year. Preferably, such durable hydrogel microparticles, when implanted, do not disintegrate under normal physiological (also referred to as normal in vivo clearance of foreign substances via phagocytosis, degradation, adsorption, etc.) conditions for at least 3 months, more preferably at least 6 months.

[0017] In either case, the hydrogel microparticles may be characterized as microspheres, microbeads, or hydrogel microparticles. The hydrogel microparticles are in the form of individual free-standing bodies that include a three-dimensional hydrogel matrix and cells suspended, entrapped, or encapsulated in the hydrogel matrix. A sustained release composition for implantation in a subject is also described herein. The sustained release composition comprises a plurality of three-dimensional hydrogel microparticles suspended in a pharma- ceutically acceptable delivery vehicle.

[0018] Suitable hydrogel precursor compounds used to form the microparticles include hydrogel-forming polymers, oligomers, and / or monomers, and as such are capable of forming a crosslinked or network-type structure or matrix (i.e., "hydrogel") through crosslinking, and fluids and cells may be retained, suspended, entrapped, and / or encapsulated within the interstitial spaces or pores of the resulting gelled structure or matrix body. The hydrogel precursor compounds used in the present invention are preferably non-alginate-based hydrogel precursor compounds. That is, the hydrogel precursor solution is preferably substantially free of alginate-based compounds, i.e., compounds based on alginate esters, alginic acid, or salts or derivatives thereof. The term "substantially free," as used herein, means that the component has not been intentionally added to the composition, even if incidental impurities or residual / traces from manufacturing processes may occur. In such embodiments, the hydrogel precursor solution composition contains less than about 0.05 wt. %, preferably less than about 0.01 wt. %, and more preferably about 0 wt. % of such components, based on the total weight of the solution taken as 100 wt. %.

[0019] The hydrogel microparticles can be advantageously prepared from a variety of slow gelling polymer precursors, such as hyaluronic acid (HA), polyethylene glycol diacrylate (PEGDA), polyethylene glycol maleimide (PEGMAL), hyperbranched PEG, hyaluronan, fibrin, chitosan, collagen, heparin, polylactic acid (PLA), poly(L-lactic acid) (PLLA), polylactic-co-glycolic acid (PLGA), polycaprolactone (PCL), polyvinyl alcohol (PVA), polyvinylpyrrolidone (PVP), methacrylic acid (MAA), 2-hydroxyethyl methacrylate (HEMA), polyacrylamide (PAM) extracellular matrix, and their functionalized (e.g., acrylated, methacrylated, thiolated, etc.) and hyperbranched (e.g., 4-armed, 8-armed) variants. However, any crosslinkable hydrogel precursor compound is suitable for use in the present invention, with preferred compounds being biocompatible non-alginic homopolymers or copolymers, particularly non-alginic block copolymers, as well as other types of crosslinkable monomers and / or oligomers. Typically, the amount of polymer precursor contained in the precursor solution will be less than 50 w / w%. This amount ranges from about 1 w / w% to about 50 w / w% for high molecular weight (500 kDa or more, preferably 100 kDa or more) polysubstituted polymers such as hyaluronic acid, and from about 5 w / w% to about 50 w / w% for lower molecular weight polymers (less than 100 kDa, preferably 50 kDa or less), such as PEGDA and PEGMAL.

[0020] The crosslinking properties of the microparticles can be tuned by adjusting the molecular weight of the precursor compounds as well as the crosslinking agent, crosslinking conditions, and crosslinking process selected. For example, in some embodiments, a "tighter" or faster crosslinking may be desired to limit the ability of molecules to leach out of the droplets / cores into the surrounding environment upon crosslinking, resulting in a smoother bead surface or a more rigid gel. This can be achieved by increasing the molecular weight of the precursor species (e.g., above about 40 kDa) and / or decreasing the crosslinking time by adjusting the crosslinking chemistry and initiation method. However, increasing the molecular weight of the bifunctional precursor species (i.e., PEGDA) typically results in a more diffuse gel due to a decreased crosslinking density. Furthermore, the use of readily hydrolyzable crosslinkers can produce microparticles with low initial Q values ​​and initially rigid to facilitate cell engraftment at the implantation site. However, such crosslinkers are rapidly degraded under normal physiological conditions, resulting in a rapid release of their payload at the implantation site. Crosslinking can be accomplished by a variety of mechanisms depending on the particular hydrogel precursor compound. In one or more embodiments, the core / shell microparticles are combined, preferably in solution, with a hydrogel matrix crosslinker. The crosslinker leaches through the alginate shell into the core / shell microparticles, resulting in gelation (crosslinking) of the hydrogel precursor compounds to form a three-dimensional hydrogel matrix. The crosslinker will correspond to the hydrogel precursor compounds, but can be varied to control the rate and level of crosslinking achieved in the resulting crosslinked matrix. Typically, the amount of crosslinker used in forming the microparticles ranges from about 0.5 mM to about 30 mM, preferably from about 1 mM to about 20 mM, more preferably from about 2 mM to about 15 mM, even more preferably from about 2.5 mM to about 10 mM, and even more preferably from about 2.5 mM to about 5 mM, depending on the crosslinker and polymer system selected.

[0021] Suitable crosslinkers include photo- or thermally initiated crosslinkers, chemical crosslinkers such as acrylates, methacrylates, acrylamides, vinyl-sulfones, dithiols, etc., which will be included as part of the alginate bath. Self-crosslinking hydrogel precursors can also be used. In these embodiments, a photoinitiator such as IRGACURE® 2959 (2-hydroxy-4'-(2-hydroxyethoxy)-2-methylpropiophenone) or lithium phenyl-2,4,6-trimethylbenzoyl-phosphinate (LAP) can be included in the hydrogel precursor solution as a catalyst. The photoinitiator can also be included in the alginate bath. For chemical crosslinking systems, the crosslinker is typically provided in the alginate bath. For UV-initiated crosslinking systems, the crosslinker can be included in the alginate bath or in the hydrogel precursor solution along with the main polymer (backbone) species.

[0022] Table 1 shows examples of reactive groups with different backbone chemistries that can be used to form hydrogels with the appropriate crosslinkers. Some of these reactions are initiated by UV light, others are chemical reactions. [Table 1]

[0023] Further exemplary precursor compounds include, but are not limited to, non-alginate polysaccharides, collagen / gelatin, chitosan, agarose, etc. These precursor compounds can be branched polymers with multiple branches or can be single backbone unbranched / linear polymer chains. Additionally, these polymers can be further functionalized by attaching proteins, drugs, or enzymes to the hydrogel to modulate cell attachment, function, proliferation, or viability. A particularly preferred hydrogel precursor compound is hyaluronic acid, or a hyaluronic acid / PEG mixture. The precursor compounds can also be functionalized with various chemical moieties that are added to the precursor compound during hydrogel fabrication. These chemicals are bound to the hydrogel matrix as opposed to being encapsulated in the voids due to simple diffusion out of the microparticle. Examples include the addition of crosslinkers or cell support agents that can be functionalized to react with the hydrogel backbone. In rapidly degrading microparticles, these agents are released locally along with the cells and cell products encapsulated in the microparticle.

[0024] To reiterate, preferred hydrogels are slow-gelling hydrogels, which as used herein means hydrogels that have an insufficient gelling rate to form constructs having normal smoothness and normal spherical morphology upon contact of gel precursor droplets with a solution containing a crosslinker. Slow-gelling hydrogels are generally those that do not instantly (or nearly instantly, e.g., within 5-10 seconds) form gel droplets upon exposure to a crosslinker. Biocompatible hydrogels are also particularly preferred, depending on the intended end use of the hydrogel. As used herein, "biocompatible" means that it is not harmful to living tissues or cells encapsulated within the matrix, more specifically, it is not biologically or otherwise undesirable in that it can be administered to a subject without excessive toxicity, irritation, allergic reaction, or immunogenic reaction, and it does not cause any undesirable biological effects or interact adversely with any of the other components of the composition in which it is contained. As known to those skilled in the art, a biocompatible hydrogel is selected to minimize any adverse side effects in the subject. Additional optional components that may be included with the hydrogel precursors include extracellular matrix components such as fibronectin, laminin, collagen, and the like, including synthetic versions thereof.

[0025] Typically, the method for making the microparticles is detailed in U.S. Patent No. 9,642,814, filed June 3, 2015, which is incorporated herein by reference. The method includes providing a hydrogel precursor solution including hydrogel precursor compounds, cells to be encapsulated, and divalent cations (e.g., calcium, barium, strontium, and combinations thereof) dispersed or dissolved in a solvent system. The divalent cations are dispersed or dissolved in the solvent system along with the hydrogel precursor compounds. The divalent cations may be present in the solution at a level of about 0.025 moles / liter to about 0.25 moles / liter, based on the total volume of the solution taken as 100%.

[0026] The hydrogel precursor solution may also include any hydrogel crosslinkers, catalysts, additives, media, nutrients, pH buffers, density modifiers, or viscosity modifiers. The hydrogel precursor solution is combined with alginate to initiate gelation of the alginate around the hydrogel precursor solution (via an "inside-out" gelation process) to yield core / shell microparticles. Each core / shell microparticle includes an alginate shell around a liquid core, which includes the hydrogel precursor solution. As shown in FIG. 2, this typically involves dropwise addition of the (A) precursor solution to a (B) alginate bath, for example by generating / extruding droplets of the precursor solution and dripping or spraying them into the alginate bath. The amount of alginate in the solution may vary, but may range from about 0.1% w / v to about 2.0% w / v, based on the total volume of the solution taken as 100%. Typically, the viscosity of the alginic acid solution should be less than that of the hydrogel precursor solution. The viscosity of the alginic acid solution depends on the alginic acid concentration and the average molecular weight of the alginic acid polymer (i.e., the length of the alginic acid molecule or the number of monomer units in the chain), with the longer the chain, the higher the viscosity at the same concentration. In one or more embodiments, the viscosity of the alginic acid solution ranges from about 1 to about 20 cP, preferably from about 1 to about 4 cP, at room temperature (about 20 to 25° C.). More specifically, the ratio of the viscosity of the hydrogel precursor solution to the viscosity of the alginic acid solution should be greater than 1 at room temperature. In one more embodiments, the ratio of the viscosity of the hydrogel precursor solution to the viscosity of the alginic acid solution is about 1:1 to about 1000:1. In one or more embodiments, the ratio of the viscosities of the hydrogel precursors is about 20:1. In one or more embodiments, the viscosity of the hydrogel precursor solution is about 1 to about 500 cP, with about 40 to about 100 cP being particularly preferred at room temperature. The pH of the alginate bath should be in the range of about 6.2 to about 7.8, and preferably in the range of about 6.6 to about 7.4.

[0027] As shown in (B) and (C), an alginate shell forms from the inside and thickens around the droplet as cations leach out of the droplet of precursor solution. In other words, the presence of cations in the droplet causes the alginate in the bath to aggregate on the surface and form crosslinks around the droplet. Gelation of the hydrogel precursor compounds in the liquid core is then initiated, such as by crosslinking and / or polymerization, to yield core / shell crosslinked microparticles. Each core / shell crosslinked microparticle includes an alginate shell and a gelling core, which includes a crosslinked three-dimensional hydrogel matrix and encapsulated cells. Crosslinking can be chemically introduced, thermally introduced, or photoinitiated, depending on the particular precursor solution prepared. For example, a hydrogel crosslinker can be included in the alginate bath and allowed to diffuse through the alginate shell to crosslink the hydrogel microparticles in the core. Alternatively, crosslinking can be initiated in the hydrogel microparticle core by exposing the core / shell microparticles to an ultraviolet light source. UV exposure times range from about 1 minute to about 10 minutes, preferably from about 2 minutes to about 8 minutes, and even more preferably from about 2.5 minutes to about 5 minutes. The wavelength of UV exposure depends on the photoinitiator and / or polymer system selected, but typically ranges from about 100 nm to about 400 nm, preferably from about 315 nm to about 400 nm, and more preferably about 365 nm.

[0028] Unlike the ionic bonds of alginate-based microparticles, the microparticles of the present invention include covalent crosslinking of hydrogel precursor compounds, as shown in FIG. 1. In the example of PEGDA, the PEG backbone (black line) is shown with diacrylate chemistry at the end, and reactive sites are shown with double red lines. The PEGDA molecules crosslink with each other to produce the final hydrogel formulation. In contrast, the MeHA backbone (blue line) with reactive groups (double red lines) shown dispersed on the chain is covalently crosslinked with PEGDA (FIG. 1). It will thus be appreciated that the precursor compounds undergo a chemical transformation in the final product via a crosslinking reaction to obtain a crosslinked hydrogel matrix. For example, in one or more embodiments, the hydrogel matrix essentially consists of (or may even consist of) a crosslinked network of PEG and HA polymers containing thioether and ester crosslinks (e.g., in the case of thiolated HA). However, it will be appreciated that the matrix may contain trace amounts of unreacted precursor compounds, functional groups, etc. remaining in the network. Other examples of possible hydrogel matrices and crosslinks are shown in Table 1. This list is exemplary and not exhaustive. Examples of scaffold molecules that can be used in the chemistries listed in Table 1 include chitosan, agarose, chondroitin sulfate, or combinations of these molecules. The bonds formed between the scaffolds include thioester+ester chemical bonds, thioester+ester and methyl on the beta carbon chemical bonds, thioethersulfone chemical bonds, or thioethersuccinimide chemical bonds. However, other chemical bonds may be used. Photo (free radical) bonds may include ester+ether or amide+ether. It will be appreciated that the combinations available are complex, but the possible combinations are nearly limitless.

[0029] After the core is gelled, the alginate shell is removed (e.g., by a chelating agent and / or mechanical agitation such as sonication) to yield free-standing hydrogel microparticles or microbeads with cells encapsulated therein, as shown in (D). In other words, the alginate shell is not part of the final product and is always removed prior to use of the microparticles. The resulting hydrogel microparticles can be recovered from the solution using a mesh screen or other device and, if desired, rinsed or suspended in culture medium or appropriate nutrients. Again, the resulting degradable hydrogel microparticles are preferably substantially free of alginate.

[0030] Using this method, cells, cell clusters, tissues, combinations thereof, and fragments thereof can be encapsulated / entrapped within the hydrogel microparticles. Cells can be naturally occurring cells, i.e., freshly isolated primary cells, cultured cells, i.e., expanded cells, established cell lines, differentiated or undifferentiated cells, engineered or genetically modified cells, etc. Non-limiting examples of cells and tissues include pancreatic islets, pancreatic islet clusters, hepatocytes, stem cells, and related cells and tissues, as well as 3-dimensional cell clusters for tissue engineering or cell-based therapy, such as endocrine cells, stem cell clusters, thyroid clusters, adrenal clusters, pituitary clusters, etc. Combined cell types and / or tissues can also be used within the microparticles (e.g., primary cell types combined with stem cells). The microparticles can include cell type-dependent cell media components that support cell maintenance and growth within the hydrogel matrix. Exemplary media components include one or more of serum, nicotinamide, antibiotics (penicillin, amphotericin B, streptomycin, gentamicin sulfate), amino acids (alanine, arginine, aspartic acid, glutamine, etc.), pH buffers (sodium bicarbonate), inorganic salts (sources of sodium, potassium, magnesium, and calcium ions), carbon sources (glucose, galactose, fructose, maltose, sodium pyruvate), proteins and peptides (albumin, transferrin, fibronectin, activin, insulin), fatty acids and lipids, vitamins (A, D, E, K, B, nicotinamide), minerals and trace elements (zinc, copper, selenium), hormones, growth factors, and the like. Complete media, basal media, and media components are commercially available and include, but are not limited to, Eagle's Minimum Essential Medium (EMEM), Dulbecco's Modified Eagle's Medium (DMEM), RPMI-1640, Ham's Nutrient Mixture, Iscove's Modified Dulbecco's Medium (IMDM), CMRL-1066, and the like.

[0031] The resulting hydrogel matrix is ​​characterized as a semi-rigid network that is permeable to liquids and gases, but does not flow and retains its integrity at steady state. Hydrogels can be considered solids and can also be described as having a viscosity approaching infinity. The hydrogel matrix is ​​three-dimensional and self-supporting. The term "self-supporting" means that once formed, the hydrogel matrix will retain its shape without an external supporting structure and is not prone to deformation by its own mere internal force or weight. A self-supporting body is not flexible, permanently deformable, or flowable like a jelly, putty, or paste, but is elastic such that the matrix body may yield or deform temporarily under force. In other words, a self-supporting body will recoil or spring back to its original shape after slight compression and / or flexing, although it is understood that the hydrogel matrix may crack, break, or shear (without recovering) under sufficient external pressure or force.

[0032] The hydrogel microparticles are matrix-type capsules that retain the loading throughout the bead, rather than having a separate shell as in core-shell capsules. As mentioned above, the hydrogel microparticles are also self-supporting. In one or more embodiments, the resulting microparticles are substantially spherical in morphology. The particle size is advantageously highly customizable depending on the capabilities of the selected droplet generator. In one or more embodiments, the resulting hydrogel microspheres or hydrogel microparticles have an average maximum surface-to-surface dimension (i.e., the diameter in the case of spherical or ellipsoidal microspheres) of greater than 30 μm, and may be as large as 300 μm. In one or more embodiments, the resulting hydrogel microspheres or hydrogel microparticles have an average maximum surface-to-surface dimension of less than about 5 mm. Preferably, the resulting hydrogel microspheres or hydrogel microparticles have an average maximum surface-to-surface dimension of less than about 2 mm, more preferably in the range of about 30 μm to about 2 mm, even more preferably in the range of about 50 μm to about 1.5 mm, more preferably in the range of about 150 μm to about 1.5 mm, and even more preferably in the range of about 300 μm to about 1.4 mm. In some cases, smaller microparticles can be formed, ranging in size from about 30 μm to about 750 μm or 500 μm. For ease of reference, this cross-sectional dimension will simply be referred to as the "size" of the microparticle. In any event, the hydrogel microparticles of the present invention are not nanosized and are not to be considered nanoparticles or any other type of nanocrystalline shape.

[0033] The hydrogel microparticles are particularly suited for localized therapeutic treatment of acute or chronic conditions for which short-term treatment protocols may be used to alleviate them, including localized delivery (e.g., direct injection) of cells, cell masses, and / or tissues for bone or cartilage repair (e.g., joints) and for general protection of the implanted therapeutic cells, cell masses, and / or tissues from the host immune system. The microparticles provide short-term protection (e.g., days, weeks, up to 3 months) for the encapsulated cells, cell masses, and / or tissues from the immune system, releasing a therapeutic effect until the hydrogel breaks down and is washed away by normal physiological processes. Systemic therapy is also contemplated. In one or more embodiments, the microparticles are not intended to deliver small molecule therapeutic agents and preferably do not include such agents, unless such agents are included in the microparticles as cell-supporting active agents (e.g., for the success of cells or grafts, such as immunosuppressants) and not as active or therapeutic agents themselves.

[0034] In the therapeutic method, the microparticles are suspended or dispersed in a delivery vehicle suitable for administration to a subject. Exemplary delivery vehicles include a biocompatible suspension, viscous solution, putty, pasta, or gel in which the microparticles are distributed, preferably including nutrients and ingredients that support the maintenance and growth of the cells, cell mass, and / or tissue to be administered. Cell culture media, such as those described above, which may be further supplemented based on the type of cells and tissue, are preferred vehicles. Buffers such as saline may also be used as delivery vehicles. The method typically includes (or consists of) administering a therapeutically effective amount of the resulting microparticle composition to a patient, for example, locally to the site of inflammation, injury, arthritis, degeneration, etc. of the patient. Administration typically includes injecting the microparticle composition directly to or near the site of inflammation, injury, arthritis, degeneration, etc. The sustained release microparticle composition is advantageous because it confines the administered cells, cell mass, and / or tissue to a localized area and slowly releases the cells, cell mass, and / or tissue and / or their cellular components or cell products to the localized area for a therapeutically effective period, thereby improving or reducing the severity of the condition in the patient. As used herein, the term "therapeutically effective" refers to an amount and / or period that induces a biological or medical response in a tissue, system, animal, or human that is the subject of a researcher or clinician's search, particularly inducing some desired therapeutic effect. For example, in one or more embodiments, the therapeutically effective amount and therapeutically effective period are those that reduce inflammation and initiate or promote healing at the site of inflammation, injury, arthritis, degeneration, and the like. It will be understood by those skilled in the art that an amount or period may be considered therapeutically effective even if the condition is not completely eradicated but is only partially improved.

[0035] If necessary, the treatment may be repeated with additional injections or infusions. Those skilled in the art will appreciate that treatment protocols may be modified depending on the particular inflammation, injury, arthritis, degeneration, condition, or healing state, as well as the preferences of the physician, veterinarian, or researcher. For example, neuronal repair, such as for repairing spinal cord injury or degenerative disease, may be treatable by injecting microencapsulated stem cells into the epidural space, i.e., epidurally into the affected area. Degradable hydrogel microparticles may be designed to immediately release stem cells into the area to allow the stem cells to directly interact with the environment, or degradable microparticles designed to last longer (e.g., about 1-3 months) may microencapsulate stem cells and release positive growth factors into the area while slowly degrading to release the cells for destruction and degradation until nothing remains. Similarly, cardiac repair following a heart attack may be improved by localized release of cells or growth factors from cells, cell masses, and / or tissues that have been microencapsulated and placed in the injured area. Another example includes the treatment of non-functioning glands with hormone therapy, such as restoring thyroid function when scar tissue is present following removal or resection of the thyroid gland by placing microencapsulated thyroid cells, cell masses, and / or tissue in the neck or nearby / adjacent sites. Further examples of durable hydrogel microparticles include the localized delivery of neurotransmitters involved in neuroendocrine disorders, delivery of bone promoting factors delivered near non-healing fractures, systemic release of molecules such as atrial natriuretic peptide to treat congestive heart failure, release of clotting factors in the treatment of hemophilia, or encapsulation of engineered or transgenic cells that produce various biologically active substances.

[0036] In one or more embodiments, "empty" hydrogel microparticles can be administered as part of a joint lubrication treatment. In other words, the presence of the hydrogel itself provides therapeutic relief to the patient without the need for cells or any other active agents within the particle itself.

[0037] A particular advantage of the compositions and microparticles of the present invention is their improved "stickiness" when implanted, as compared to alginate and other types of microparticles. That is, the microparticles tend to adhere or stick to tissue at the site of implantation in vivo, as shown in the photographs in the examples below. This further enhances the efficacy of the treatment by maintaining the local release of cells, cell mass, and / or tissue and their cell products at the implantation site. Furthermore, the microparticles do not induce inflammatory and / or foreign body reactions, such as lymphocytes and collagen ring formation, at the implantation site.

[0038] Further advantages of various embodiments of the present invention will be apparent to those skilled in the art upon review of the disclosure herein and the examples below. It will be understood that the various embodiments described herein are not necessarily mutually exclusive, unless otherwise specifically stated herein. For example, features described or depicted in one embodiment may be included in other embodiments, but are not necessarily included therein. That is, the present invention encompasses various combinations and / or integrations of the specific embodiments described herein.

[0039] As used herein, the term "and / or," when used in a list of two or more items, means that any one of the listed items may be employed alone, or any combination of two or more of the listed items may be employed. For example, if a composition is described as containing or not containing components A, B, and / or C, the composition may contain or not contain A alone; B alone; C alone; a combination of A and B; a combination of A and C; a combination of B and C; or a combination of A, B, and C.

[0040] Numerical ranges are also used herein to quantify certain parameters in connection with various embodiments of the present invention. It should be understood that when a numerical range is provided, such a range should be interpreted as providing literal support for claim limitations that recite only the lower value of the range and for claim limitations that recite only the upper value of the range. For example, when a numerical range of about 10 to about 100 is disclosed, it provides literal support for claims that recite "greater than about 10" (without upper limit) and claims that recite "less than about 100" (without lower limit). The present invention is as follows. [1] A non-alginate based hydrogel microparticle for the localized delivery and sustained release of therapeutic cells and / or tissues at an implantation site, said microparticles comprising a three-dimensional matrix of a covalently crosslinked non-alginate based polymeric compound and a therapeutically effective amount of cells and / or tissues encapsulated therein, said cells having a viability of at least about 50%, and said microparticles having a size of greater than about 30 μm. [2] The microparticles described in [1] above, having a size of more than 300 μm. [3] The microparticle according to [1] above, wherein the polymer compound is selected from slow gelling polymer precursors selected from the group consisting of branched or unbranched hyaluronic acid, branched or unbranched functionalized hyaluronic acid, branched or unbranched functionalized polyethylene glycol, hyaluronan, fibrin, chitosan, collagen, polylactic acid, poly(L-lactic acid), polylactic-co-glycolic acid, polycaprolactone, polyvinyl alcohol, and combinations thereof. [4] The microparticles described in [1] above, wherein the matrix further comprises a crosslinker crosslinked with the non-alginic acid polymer compound selected from the group consisting of dithiothreitol, branched or unbranched functionalized polyethylene glycol, dithiols, ethylene glycol bis-mercaptoacetate, and combinations thereof. [5] The microparticle according to [4] above, wherein the functionalized polyethylene glycol is selected from the group consisting of polyethylene glycol dithiol, polyethylene glycol diacrylate, polyethylene glycol divinyl sulfone, polyethylene glycol dimaleimide, and combinations thereof. [6] The microparticles described in [1] above, wherein the matrix consists essentially of a functionalized hyaluronic acid precursor compound and a functionalized polyethylene glycol precursor compound, which are covalently crosslinked. [7] The microparticles according to [1] above, wherein the matrix consists essentially of a self-crosslinked functionalized polyethylene glycol precursor compound. [8] The microparticles described in [1] above, wherein the matrix essentially consists of self-crosslinked functionalized hyaluronic acid precursor compounds. [9] The microparticle of [1] above, wherein the matrix consists essentially of first and second functionalized polyethylene glycol precursor compounds covalently crosslinked, the first and second functionalized polyethylene glycol precursor compounds being different from each other.

[10] The microparticles described in [1] above, wherein the matrix essentially consists of a methacrylated hyaluronic acid precursor compound and a polyethylene glycol diacrylate precursor compound crosslinked by a covalent bond.

[11] The microparticles described in [1] above, wherein the matrix essentially consists of a thiolated hyaluronic acid precursor compound and a polyethylene glycol diacrylate precursor compound crosslinked by a covalent bond.

[12] The microparticle according to [1] above, wherein the matrix essentially consists of a polyethylene glycol maleimide precursor compound and a polyethylene glycol dithiol precursor compound crosslinked by a covalent bond.

[13] The microparticle of [1] above, wherein the matrix essentially consists of a polyethylene glycol vinyl sulfone precursor compound and a dithiothreitol or ethylene glycol bis-mercaptoacetate precursor compound, which are covalently crosslinked.

[14] The microparticle according to [1] above, wherein the microparticle contains the cells or tissue in up to 50% of its volume.

[15] The microparticles according to [1] above, which have a storage stability in PBS at 37°C for less than 6 months.

[16] The microparticles described in [1] above, which have a storage stability in PBS at 37°C for more than 6 months.

[17] The microparticle according to the above [1], which is a degradable microparticle that decomposes in less than 3 months when transplanted in vivo.

[18] The microparticles according to the above item [1], which are durable microparticles that do not decompose for at least three months when implanted in a living body.

[19] The microparticle according to [1] above, further comprising a cell culture medium component.

[20] The microparticle described in [1] above, wherein the cells are non-proliferating cells or proliferating cells.

[21] The microparticle of [1] above, further comprising, in addition to cells, secreted cell products such as signaling molecules, cytokines, chemokines, therapeutic proteins, hormones, vesicles, antibodies, viruses, exosomes, and the like.

[22] The microparticle described in [1] above, wherein the cell is a genetically engineered cell or a transgenic cell.

[23] The microparticle described in [1] above, wherein the cells are a cell mass consisting of a plurality of aggregated cells.

[24] The microparticle described in

[23] above, wherein the cell mass contains two or more different types of cells.

[25] The microparticles according to [1] above, wherein the matrix is ​​homogeneous and comprises a polymer backbone crosslinked by one type of covalent bond.

[26] The microparticles of [1] above, wherein the matrix is ​​heterogeneous, comprising a mixture of two or more polymer precursors, such as a combination of a high molecular weight polymer precursor and a low molecular weight polymer precursor, and / or a mixture of two or more crosslinkers.

[27] The microparticle according to [1] above, having a size of about 50 μm to about 5 mm.

[28] A composition for localized delivery and sustained release of therapeutic cells and / or tissues and / or cell products to a transplantation site, comprising a plurality of hydrogel microparticles according to any one of [1] to

[27] above dispersed or suspended in a biocompatible delivery vehicle.

[29] The composition described in

[28] above, further comprising a plurality of hydrogel microparticles comprising a three-dimensional matrix of a non-alginate-based polymer compound crosslinked by covalent bonds, having a size greater than about 30 μm, and essentially free of cells or tissue.

[30] The composition described in

[28] above, wherein the plurality of hydrogel microparticles comprises a mixture of degradable hydrogel microparticles and durable hydrogel microparticles dispersed or suspended in the biocompatible delivery vehicle.

[31] The composition described in

[30] above, wherein the durable hydrogel microparticles contain insulin-producing cells and / or tissues and the degradable hydrogel microparticles contain stem cells.

[32] The composition described in

[28] above, wherein the plurality of hydrogel microparticles comprises a plurality of first hydrogel microparticles comprising a first cell type and / or tissue type, and a plurality of second hydrogel microparticles comprising a second cell type and / or tissue type different from the first cell type and / or tissue type.

[33] The composition of

[32] above, wherein the first cell and / or tissue type comprises insulin-producing cells and / or tissues and the second cell and / or tissue type comprises stem cells, or glucagon-producing cells and / or tissues.

[34] The composition described in

[28] above, wherein the plurality of hydrogel microparticles comprises a plurality of first hydrogel microparticles having a first size and a plurality of second hydrogel microparticles having a second size different from the first size.

[35] The composition described in

[28] above, wherein the vehicle is a biocompatible liquid suspension or gel.

[36] The composition described in

[28] above, wherein the medium is a solution or gel of a cell culture medium.

[37] The composition according to

[28] above, which is injectable or implantable via a catheter.

[38] The composition described in

[28] above, comprising a therapeutically effective amount of the microparticles.

[39] The composition according to

[28] above, which is in the form of a putty, paste, or gel.

[40] The composition described in

[28] above, which is in the form of a liquid suspension.

[41] A method for locally delivering and sustained releasing therapeutic cells and / or tissues to a transplantation site, comprising administering to a subject a plurality of microparticles described in any one of [1] to

[27] above or a composition described in any one of

[28] to

[40] above to the transplantation site.

[42] The method of

[41] above, wherein said administering comprises injecting said composition at or near the implantation site.

[43] The method according to

[41] above, wherein the injection is an intra-articular injection.

[44] The method according to

[41] above, wherein the transplantation site is an inflamed area, an injured area, an arthritic area, a degenerated area, etc. of the subject.

[45] The method according to

[41] above, wherein the transplant site is a joint.

[46] The method according to

[41] above, wherein the implantation site is an area near a gland and the microparticles release cellular products, such as hormones, near the gland.

[47] The method according to

[41] above, wherein the implantation site is within the intracapsular cavity, i.e., within a physiologically contained space such as within the pericardium or renal capsule.

[48] ​​The method according to

[41] above, wherein the transplantation site is the subject's omentum or peritoneum.

[49] The method according to

[41] above, wherein the transplantation site is a tumor site or a site adjacent to the tumor site.

[50] The method according to

[41] above, wherein the transplantation site is a site after tumor resection or a site adjacent to the site.

[51] The method according to

[41] above, wherein the transplant site is a septic site.

[52] The method according to

[41] above, wherein the microparticles are biodegraded within about 3 months after administration.

[53] The method according to

[41] above, wherein the microparticles are not biodegradable for at least six months after administration.

[54] The method according to

[41] above, wherein the implantation site is a joint and the microparticles are mechanically degraded within about 3 months after administration.

[55] The method according to

[41] above, wherein the implantation site is a joint and the microparticles do not mechanically degrade for at least 6 months after administration.

[56] The method described in

[41] above, wherein the hydrogel microparticles adhere to the tissue at the transplantation site, thereby maintaining the transplanted cells at the transplantation site during the treatment period.

[57] The method of

[41] above, wherein cells and / or tissues and / or cell products are released from the microparticles through the pores of the microparticles and / or via biodegradation of the microparticles to a localized area at the implantation site, thereby alleviating or reducing the severity of the condition in the subject.

[58] The method according to

[41] above, wherein the microparticles are biodegradable within about 5 days after administration.

[59] The method according to

[41] above, wherein the microparticles are biodegradable within about 24 hours after administration.

[60] The method according to

[41] above, wherein the microparticles are biodegraded within about one week after administration.

[61] The method according to

[41] above, wherein the microparticles are biodegraded within about one month after the administration.

[62] The method according to

[41] above, wherein the microparticles are biodegradable within less than about 6 months after administration.

[63] The method according to

[41] above, wherein the microparticles are biodegraded after administration, and the biodegradation comprises hydrolysis of the polymer backbone.

[64] The method according to

[41] above, wherein the microparticles are biodegraded after administration, and the biodegradation comprises hydrolysis of the crosslinker.

[65] The method according to

[41] above, wherein the microparticles biodegrade after administration, and the degradation rate is adjustable by varying a parameter selected from the group consisting of the molecular weight of the polymer precursor, the molecular weight of the crosslinker, the ratio of the crosslinker to the polymer precursor, hydrolysis of the crosslinker, crosslinking kinetics, ultraviolet light exposure time, and combinations thereof.

[66] A method for reducing pain and / or providing viscosupplementation in a joint, comprising introducing into the joint a plurality of hydrogel microparticles comprising a three-dimensional matrix of a covalently crosslinked non-alginate based polymeric compound, the microparticles having a size greater than about 30 μm, and being essentially free of cells or tissue. EXAMPLES

[0041] The following examples describe methods according to the present invention, however, it should be understood that these examples are provided by way of illustration and that no one should be construed as a limitation on the overall scope of the invention.

[0042] The HA formulations used in each example are shown to degrade in less than 3 months. In contrast, the PEGDA formulations are more durable. The properties of the products thus produced are compared by comparing degradable and durable microparticles. Such comparisons include physical and chemical differences, in vitro and in vivo stability, cell morphology, function, and migration within the final microparticles, and the ability of the final microparticles to attach to tissue in vivo, thus localizing their application within the body.

[0043] Example 1 Physical and chemical differences between durable and degradable hydrogel microparticle end products Introduction The concept of cell encapsulation was first popularized in 1980 by Lim and Sun, who showed that pancreatic islets embedded in alginate hydrogel microparticles could reverse diabetes in rats without the need for immunosuppression, albeit for only a few weeks. This initial success was followed by studies aimed at understanding and improving the process. Although the field has made great strides over time, it is acknowledged that alginate microspheres still present issues related to their long-term bioincompatibility. Despite such limitations, alginate has remained the clear material of choice for cell encapsulation, due to its unique, nearly instantaneous crosslinking kinetics, which allows for the easy production of affordable, injectable microparticles.

[0044] Conversely, other hydrogels have slower cross-linking kinetics and can typically only be fabricated as large macroscopic structures. Several protocols have been developed to fabricate microspheres using alternative hydrogels such as polyethylene glycol, agarose, chitosan, or hyaluronic acid. However, these approaches are based on oil emulsion technology, which is generally not suitable for cell encapsulation and implantation due to its reliance on non-aqueous solvents and poor process control.

[0045] As cell encapsulation applications continue to grow in number and sophistication, it would no doubt be useful to be able to produce easily injectable biocompatible microparticles using advanced biomaterials. Herein, we have explored a novel method for producing hydrogel microparticles, termed Core-Shell Spherification (CSS), described in U.S. Patent No. 9,642,814, filed June 3, 2015, which is incorporated herein by reference. This method is strategically designed for use with hydrogels that have significantly slower gelation rates compared to alginate, allowing for the production of hydrogel microparticles with a variety of chemical, physical, and bioactive properties. Furthermore, the method does not use oil emulsion technology and was developed for standard GMP-compatible equipment and materials to better facilitate accessibility, scale, and regulatory compliance. We demonstrate microparticle production by CSS using two commonly used hydrogels: hyaluronic acid (HA) and polyethylene glycol diacrylate (PEGDA).

[0046] method Cytotoxicity of the CSS crosslinking method The cytotoxicity of calcium, a component of the CSS method, was tested by exposing canine islets to a solution containing 100 or 200 mM calcium chloride, 10 mM HEPES buffer, and 20% PEGDA (molecular weight 3,400 Da) (Laysan Bio, Inc.) for 5, 10, or 15 minutes. The effect of calcium alone was also evaluated by exposing a group to 100 mM calcium chloride without PEGDA. Islets were suspended in the test solution at approximately 5,000 IEQ / mL to mimic the high cell loading density associated with the encapsulation method. Following calcium exposure, islets were washed twice with CMRL1066 islet supplement medium and incubated at 37°C and 5% CO2 for 3 hours prior to evaluation.

[0047] Dead cells were identified using propidium iodine staining and fluorescence microscopy. Fluorescence microscopy images were captured using a Cytation5 imaging multimode reader (Biotek Instruments, Inc, 531 / 647 nm ex / em). Cell death was quantified by calculating the percentage of red (dead cell) pixels relative to total islet pixels using Adobe Photoshop. Twenty-five individual islets per group were analyzed and differentiated from non-islet tissue by dithizone staining. Results are reported as the mean percentage of live cells and normalized to islet viability in the matched untreated control group (i.e., islets from the same batch in medium).

[0048] In the case of PEGDA and methacrylated hyaluronic acid (MeHA) microparticles, photocrosslinking was utilized. Thus, studies were first performed to investigate the toxicity of ultraviolet (UV) light and photoinitiators. Canine islets were suspended in Dulbecco's phosphate buffered saline (DPBS) containing 0% (w / v), 0.025% (w / v), or 0.05% (w / v) IRGACURE® 2959 photoinitiator. Approximately 50 IEQ was loaded into each well of a 24-well plate and exposed to long-wave ultraviolet (UV) light for 3, 5, or 10 minutes. Plates were irradiated at approximately 40 mW / cm according to the manufacturer's data (Ubitron International). 2Exposure was performed using a PortaRay 400 UV lamp in low power mode at a distance of 6 inches, equivalent to 100 s. After exposure, islets were washed twice with CMRL1066 islet supplement medium and incubated at 37°C, 5% CO2 for 3 hours before evaluation. Cell death was assessed by propidium iodide staining and quantified as previously described.

[0049] Preparation of MeHA hydrogel microparticles After determining a non-toxic level of crosslinker, microparticle production was initiated. A schematic diagram of the CSS method for hydrogel microparticle production is shown in Figure 2. First, MeHA was synthesized by reacting HA (molecular weight 1 MDa, Lifecore Biomedical) with a 50-fold molar excess of glycidyl methacrylate (Sigma) in the presence of triethylamine and tert-butylammonium bromide (Sigma) in a 50:50 water:DMSO mixture for 5 days. MeHA was then dialyzed against deionized (DI) water for 2 days and then lyophilized. 1 The degree of methacrylation was determined by calculating the ratio of the relative peak area of ​​methacrylic acid protons to methyl protons using 1 H NMR (Avance AV-III 500, Bruker), and was found to be 54-72%.

[0050] Methacrylated HA:PEGDA polymer mixture (molecular weight 3.4 kDa, Lysun Bio) was prepared at 2.5%:1% (w / w) in a custom buffer solution containing 100 mM calcium chloride, 15 mM HEPES, and 0.05% (w / v) IRGACURE® 2959, plus OptiPrep (CosmoBioUSA, Inc.). In this embodiment, a low concentration of PEGDA was added as a crosslinker to the MeHA solution to increase the crosslink density of the hydrogel. The solution was passed through a 0.22 μm filter. The viscosity of the precursor solution was measured at room temperature using a Cannon-Manning Semi-Micro calibrated glass capillary viscometer.

[0051] The precursors were extruded via an automated droplet generator into a stirred bath of 0.15% (w / v) sodium alginate (Protanal LF 10 / 60, FMC Corp.) containing 300 mM mannitol, 0.1% Tween 20, and adjusted to pH 7.6 with a custom 15 mM HEPES buffer. In the case of the MeHA precursor, 0.05% (w / v) IRGACURE® 2959 initiator was also added to the alginate bath to maintain equilibrium of IRGACURE® between the droplets and the alginate bath and to minimize leaching of IRGACURE® from the droplets after addition to the bath (which would reduce cross-linking). Droplet generation was performed using a Buchi 395-Pro ​​encapsulation device (Buchi Corporation, New Castle, DE) equipped with an air jet nozzle system and a 400 μm diameter internal liquid nozzle within a 1.5 mm coaxial air nozzle. Droplets were extruded using compressed nitrogen. The MeHA droplets formed core-shell constructs, each with an alginate shell encapsulating a hydrogel precursor solution / droplet core. The hydrogel precursor solution / droplet core was irradiated with long-wave UV light to initiate free radical photocrosslinking of MeHA and PEGDA. The irradiance at the center of the alginate bath was approximately 40 mW / cm. 2 The irradiation was carried out with a 300 nm fluoride (PortaRay 400, Ubitron International). Irradiation was performed continuously during and for 1 min after extrusion of the precursor solution to ensure complete cross-linking of the hydrogel cores in the core-shell constructs. The core-shell constructs were then rinsed with 25 mM citrate buffer in DPBS for 5 min with gentle agitation to dissolve the alginate shell. The resulting microparticles were collected using a steel mesh screen and suspended in DPBS. A second rinse with 50 mM citrate for 5 min was performed to complete dissolution and remove the alginate shell.

[0052] Preparation of ThHA hydrogel microparticles A ThHA precursor was prepared by dissolving thiolated HA (HyStem, Biotime Inc) at 1.2% (w / w) in a custom buffer containing 100 mM calcium chloride and OptiPrep (Cosmo Bio USA) to adjust the solution density. The pH of the precursor solution was adjusted to approximately 7.0 with a custom 15 mM HEPES buffer to suppress disulfide cross-linking between ThHA macromers. The viscosity of the solution was measured as above. Core-shell constructs were fabricated as above, except that cross-linking was achieved via chemical cross-linking (vs. free radical photocross-linking for MeHA microparticles). The terminal acrylic acid groups of the PEGDA molecules are bonded to the thiol groups on the ThHA via a Michael addition reaction. Briefly, the precursor solution was contacted with an alginate bath to produce a core-shell construct with a ThHA precursor core and an alginate shell. The alginate bath also contained 0.4% PEGDA (molecular weight 3.4 kDa, Lysun Bio) to act as a crosslinker and was gently stirred for 5 min. Both the alginate bath and the hydrogel precursor could contain a crosslinker (e.g., PEGDA), but in this protocol, it is preferred that only the alginate bath contains it. Crosslinking proceeded by diffusion of the smaller sized 3.4 kDa PEGDA molecules through the alginate shell into the core. The alginate bath was then diluted by half with DPBS, lowering the solution pH to 7.4. Stirring was continued for an additional 30 min to crosslink the HA precursor within the core. The core-shell constructs were collected and the alginate shell was removed to obtain microparticles, as described above.

[0053] Preparation of PEGDA hydrogel microparticles PEGDA hydrogel precursor solutions were prepared by dissolving 3.4 kDa PEGDA and 20 kDa PEGDA (Lysan Bio) at 18% (w / w) and 12% (w / w), respectively, in a buffer solution containing 100 mM calcium chloride, 10 mM HEPES, and 0.025% (w / v) IRGACURE® 2959. The solutions were filtered using a 0.22 μm syringe, and the viscosity was measured as above. The precursors were extruded into a stirred alginate bath similar to that for MeHA microparticles, but containing 0.025% (w / v) IRGACURE® 2959. All solutions for MeHA and PEGDA microparticle production were prepared in water that had been degassed by sonication to eliminate excess oxygen, a known photocrosslinking inhibitor. The core-shell constructs were irradiated as above to achieve crosslinking of the core. The constructs were further treated as above to dissolve the alginate shell and yield hydrogel microparticles.

[0054] Manufacturing of AHA hydrogel particles First, AHA was synthesized by reacting HA (molecular weight 1 MDa or 200 kDa, Lifecore Biomedicals) with acryloyl chloride and glycidol (TCI) in the presence of triethylamine and N-dimethylformamide (Sigma) for 5 days. Next, AHA was dialyzed against deionized (DI) water for 5 days and then lyophilized. The degree of acrylation was determined by calculating the ratio of the relative peak area of ​​the acylate proton to the methyl proton using 1H NMR (Avance AV-III 500, Bruker), which was 53-72%. Next, AHA precursor solution was prepared by dissolving acrylated HA at 4% in a custom buffer containing 100 mM calcium chloride, 15 mM HEPES buffer, and 0.05% (w / v) IRGACURE® 2959. This precursor solution was filtered using a 0.22 μm syringe, and the viscosity was measured as described above. The precursor was extruded into a stirred alginate bath as for MeHA microparticles. The core-shell constructs were irradiated as above to achieve cross-linking of the core. The alginate shell was removed using citrate buffer as above, and the AHA hydrogel microparticles were collected.

[0055] Physical properties and size distribution of hydrogel microparticles Individual images of 100 microparticles were taken using a Biotek Cytation5 multimode imaging reader. Microparticles were imaged in PBS in a multiple well plate, and images were analyzed in Adobe Photoshop to determine the average microparticle diameter and size distribution for each microparticle type (N=100 microparticles). The hydrogel microparticles were further characterized by determining the swelling ratio "Q", the ratio of swollen hydrated mass to dry mass, for each microparticle type. To reach equilibrium swelling, microparticles were immersed in excess deionized water for 24 hours prior to measurement to remove dissolved salts. Excess surface moisture was removed from the microparticles before weighing them on a pre-weighed watch glass. Microparticles were then dried overnight at 60°C and reweighed to obtain the dry mass (N=3).

[0056] Diffusion characteristics of fine particles Hydrogel microparticles were incubated overnight in 0.1 mg / mL FITC-labeled dextran (Invitrogen Molecular Probes) with molecular weights of 10 kDa, 40 kDa, 70 kDa, and 500 kDa in DPBS according to previously published procedures. Microparticles were rinsed with DPBS and imaged by laser scanning confocal microscopy to monitor the efflux of the probe (Olympus Fluoview 300). Microscopic images were taken at set time points, from 3 to 30 min after removal of the microparticles from the FITC-dextran incubation solution.

[0057] Pore ​​size calculation Empty PEGDA and MeHA microparticles were examined by environmental scanning electron microscopy (ESEM) performed on an FEI Quanta 600F ESEM instrument. Samples were immersed in water on a cooling platform with the temperature in the SEM chamber set at 6C. The chamber was then evacuated to 1000 Pa and the relative humidity was maintained at 100%. Images were taken at 10,000x magnification. Standard SEM images were also collected. For such images, samples were critical point dried or freeze dried, coated with 10 nm platinum, and imaged at 5 kV in the same ESEM instrument in high vacuum mode. SEM images were taken at 1000x magnification.

[0058] result Core-shell spherification procedure Hydrogel microparticles were fabricated by the CSS method summarized in Figure 2. In this method, a slow-setting hydrogel precursor solution (functionalized PEG or HA) is first encapsulated as a droplet within an alginate shell when applied to an alginate bath, allowing time for the functionalized PEG or functionalized HA to harden. The alginate shell formed around the core microparticle features a concentric ring-link morphology (Figure 3A). The alginate shell is then removed, leaving only the inner core hydrogel (Figure 3B). Figure 3B shows mostly spherical microparticles. However, other shapes are acceptable for the final product. Figure 4 further illustrates examples of acceptable end product features. Figure 4A shows an example of a perfectly spherical, more durable (also called persistent) PEGDA-based microparticle loaded with canine islet cells, with slower degradation properties. Figure 4B shows a similar, but elongated, durable PEGDA-based microparticle loaded with islets, with a three-dimensional ellipsoidal shape. Figure 4C shows a flat-edged, stem cell-loaded, fast-degrading (also called short-term) MeHA-based microparticle on the left, which has faster degradation properties. Figure 4D shows the same type of microparticle, but with a three-dimensional ellipsoidal shape. These microparticles, and other related shapes such as teardrop and ovoid, are all functional examples of CSS microparticle end products.

[0059] Cytotoxicity research During microparticle production, cross-linking is required for gelation of the core, such that the alginate shell remains present and maintains a 3-dimensional free-standing structure after removal of the shell. To optimize the CSS process, the cross-linking components of the microparticle production process were evaluated for their cytotoxic effects on freshly seeded cells (canine islets). Figure 5A shows the effect of calcium concentration and exposure time on the viability of canine islets. The three conditions tested were 100 mM and 200 mM calcium + PEGDA, or 100 mM calcium alone. For the shortest exposure time measurement of 5 min, no significant differences in cytotoxicity were identified in any of the groups. All groups had viability greater than 97.5% compared to the corresponding untreated control group (islets from the same batch in culture medium). However, a significant decrease in viability was seen in the 200 mM calcium group at each subsequent time point compared to the 5 min measurement. The 200 mM calcium group had significantly lower viability than the two 100 mM groups at both the 10 and 15 min time points. No differences were observed in either of the 100 mM groups throughout the 10 minutes. However, at 15 minutes, the 100 mM group with PEGDA showed a significant decrease in viability compared to the 5 and 10 minutes, and compared to the 100 mM group without PEGDA at 15 minutes. Interestingly, within the 100 mM group without PEGDA, no significant change in viability was observed at any time point, with a 97.6% viability at 15 minutes compared to the control group.

[0060] The effect of photoinitiator concentration and UV exposure time on cell viability is shown in Figure 5B. IRGACURE® 2959 concentrations of 0% (w / v), 0.025% (w / v), and 0.05% (w / v) were applied to canine islets at approximately 40 mW / cm 2Cytotoxicity was evaluated by irradiation with long-wave UV light exposure of 3, 5, and 10 minutes. Interestingly, the group without photoinitiator (IRGACURE® 2959) showed the highest degree of cytotoxicity. The viability of the 0% group was significantly lower at 5 and 10 minutes compared to the 3 minute exposure time. Furthermore, the 0% group showed significantly lower viability at 5 and 10 minutes compared to both IRGACURE®-containing groups. A significant difference between the 0.025 and 0.05% groups was only seen at 10 minutes, with viable cell rates of 94.5% and 87.2%, respectively.

[0061] Physical properties and size distribution of hydrogel microparticles To fully test this novel CSS platform, microparticles were prepared from two different starting materials: HA and PEGDA. To evaluate both chemically and photocrosslinked hydrogels using the CSS platform, the HA preparations were further divided into thiolated HA (ThHA) and methacrylated HA (MeHA), respectively. Table 2 shows the physical properties of the hydrogel formulations before and after crosslinking, as well as the average microparticle diameter and diameter range. [Table 2]

[0062] The precursor mass fraction of the gel precursor was 30% for PEGDA, while ThHA and MeHA were prepared from 1.2% and 2.5% solutions, respectively. The PEGDA precursor required a higher concentration to achieve a suitable solution viscosity to form well-formed microparticles. The gel formulations and corresponding precursor viscosities are shown in Table 2. ThHA was the only chemically crosslinked hydrogel, which required much more time (35 min) for crosslinking compared to the photocrosslinked gels (MeHA and PEGDA). MeHA microparticles had the largest average diameter, followed by PEGDA microparticles, which were then followed by ThHA microparticles, which were smaller than the other two groups. PEGDA microparticles showed the lowest equilibrium swelling ratio "Q", and the overall hydrogel was more compact. This is further illustrated in Figure 6, where the diameters of 100 representative microparticles composed of ThHA, MeHA, PEGDA, or AHA are plotted, showing good monodispersity of each microparticle, especially in the case of the HA material. Despite being produced using the same droplet generator and pre-crosslinking parameters, ThHA microparticles were significantly smaller in size than PEGDA and MeHA, and had a narrower diameter range compared to PEGDA. MeHA and AHA microparticles had the largest mean diameters among the four groups. Interestingly, islet-containing PEGDA particles were only slightly larger in mean diameter compared to empty PEGDA microparticles, but had improved monodispersity (938 μm for islet-containing PEGDA [CV: 8.4%] vs. 904 μm for empty PEGDA microparticles [CV: 15.7%]).

[0063] Diffusion characteristics of fine particles Figure 7 shows the diffusion of fluorescently labeled dextran into PEGDA, ThHA, and MeHA hydrogel microparticles. The microparticles were incubated overnight in fluorescent dextran of various molecular weights and immediately rinsed before being examined by confocal microscopy to assess dextran penetration and efflux rates as indicators of diffusivity. The 10 kDa probe was able to penetrate all microparticles. However, the PEGDA microparticles showed weaker fluorescence, especially near the center of the construct, suggesting a slower rate of probe penetration compared to the HA microparticles, both of which showed strong fluorescence. Furthermore, the fluorescence of the 10 kDa probe disappeared rapidly in the HA group and was absent or nearly absent 30 min after rinsing. In contrast, localized fluorescence was still observable in the PEGDA group at 30 min, again suggesting a higher diffusion barrier for the 10 kDa probe compared to both HA gels.

[0064] Low levels of fluorescence from the 40 kDa probe were detected at the periphery of the PEGDA microparticles, while little or no signal was observed in the center, indicating a diffusion barrier for the probe (Figure 7). The larger dextrans (70 kDa and 500 kDa) showed little or no fluorescent signal in the PEGDA microparticles, suggesting that penetration of these probes into the gel matrix was negligible over the overnight incubation period.

[0065] Both ThHA and MeHA microparticles showed strong fluorescence from all probes evaluated, suggesting minimal diffusion barriers even for larger molecules. Furthermore, the relative fluorescence signal in MeHA appeared significantly higher for all probes immediately after rinsing compared to ThHA, suggesting a higher concentration of probe and therefore a lower diffusion barrier. This was best exemplified by the 70 kDa probe, which was less loaded in ThHA microparticles but appeared to saturate in MeHA microparticles. All further studies focused on the hydrogel formulations with the highest and lowest diffusion barriers, PEGDA and MeHA, respectively.

[0066] Consideration We developed and evaluated a novel method to fabricate hydrogel microparticles for cell encapsulation and delivery. Current cell microencapsulation methods are primarily based on alginate spheres due to their fast gelation rate. Other hydrogels used for microencapsulation require oil emulsion techniques that are less favorable. Alternatively, the CSS method can be used with a wide variety of hydrogel materials, is compatible with standard commercially available GMP-compliant equipment, and has minimal cytotoxicity to pancreatic islets.

[0067] Low cytotoxicity is a critical feature in any cell encapsulation strategy. Therefore, some of the key enabling components of the CSS method were evaluated for their cytotoxic effects on islets, particularly calcium content and UV exposure. Not surprisingly, 200 mM calcium was rapidly cytotoxic. However, islets were found to tolerate 100 mM calcium for at least 10 min, even in highly concentrated and viscous PEGDA solutions. 2 Cytotoxicity of canine islets to long-wave (approximately 365 nm) UV exposure was minimal for up to 10 min. Unexpectedly, the group without photoinitiator showed the most cell death after UV exposure, which is in contrast to other similar studies. However, these other studies used UV light at 4–10 mW / cm2. 2Much lower UV intensities of 1000 nm and 1000 nm were used, and the photoinitiator concentrations used herein were also relatively low. Thus, the photoinitiator may have provided a protective effect in this experiment, possibly by selectively absorbing UV photons. Nevertheless, a concentration-dependent increase in cytotoxicity was observed in the IRGACURE® 2959-containing group, which is consistent with previously published studies.

[0068] The microparticles differ greatly in size, with MeHA beads having nearly twice the diameter of ThHA beads, corresponding to an approximately 8-fold difference in total volume per microparticle, yet they were fabricated using the same 400 μm nozzle system. The size difference between the groups was most likely the result of differences in the swelling kinetics of the post-fabrication volume. Hydrogel swelling is controlled by several variables, but is strongly correlated to the initial (i.e., pre-crosslinking) concentration of polymer and the density of crosslinks within the gel after fabrication. More specifically, solvent molecules (e.g., water) adsorbed into the gel polymer matrix exert an outward pressure, causing the gel to expand (swell), while crosslinks within the gel matrix resist the outward expansion. For example, hydrogels with high initial polymer concentration and low crosslink density are expected to expand in volume significantly after fabrication. Conversely, gels with a high ratio of crosslinks to polymer concentration tend to resist expansion after fabrication. In this study, the smaller final size of ThHA microparticles is probably a result of a combination of a low initial polymer fraction (1.2%) and a relatively high cross-linking density. An important note is that the volume swelling after preparation should not be confused with the swelling ratio "Q" (hydrated mass / dry mass). Specifically, Q describes the final gel structure after the volume swelling has reached equilibrium, and typically, the lower the Q value, the stronger and more compact the gel. That is, it is possible that a gel may undergo significant volume swelling but still have a relatively low Q value, which may be the case for PEGDA microparticles in this study. In contrast, ThHA microparticles experienced relatively little (if any) volume swelling, but had a higher Q value compared to PEGDA microparticles. Furthermore, MeHA microparticles swelled significantly and had a high Q value even with a low initial polymer concentration (3.5%), indicating a very low relative cross-linking density.

[0069] Diffusivity varied considerably among the microparticle groups. ThHA and MeHA microparticles showed greater permeability to dextran probes up to 500 kDa in size, with MeHA microparticles appearing to have the highest diffusivity of all three groups. Conversely, PEGDA particles appeared to strongly limit the diffusion of dextrans with molecular weights of 40 kDa and above. Although a variety of factors can affect the diffusivity of gels, the swelling ratio "Q" (the ratio of the equilibrium hydrated mass to the dry mass of the gel) correlates strongly with the permeability of hydrogels. PEGDA, ThHA, and MeHA microparticles had Q values ​​of 17.3, 27.7, and 105.7, respectively. Thus, the observed diffusion behavior of dextran probes in microparticles is consistent with the corresponding Q values ​​and measured surface pore sizes.

[0070] Example 2 Stability of the final products of durable and rapidly degrading hydrogel microparticles Introduction The explanation for durable versus fast degrading hydrogel microparticles relies on the stability of the microparticle's physical composition. Both in vitro and in vivo stability studies were performed to determine whether there were differences between the three formulations (PEGDA, MeHA, and AHA). While the CSS method allows for fine tuning of the hydrogels to degrade at different rates, below are exemplified two durable hydrogels that can last for months to years versus a more rapidly degrading hydrogel.

[0071] method In vitro durability testing For MeHA, PEGDA, and AHA, 1.000 grams of hydrated microparticles were collected and then added to a degradation buffer consisting of phosphate buffered saline buffered with 10 mM HEPES and 1% gentamicin. The degradation solution was maintained at pH 7.3 ± 0.10. The dissolved microparticles were stored at 37°C for 1 week, 3 weeks, and 6 weeks, then collected and dried at 60°C overnight to obtain the post-degradation dry mass. The post-degradation dry mass was compared to the microparticle dry mass on day 0 to obtain the mass loss percentage. Additionally, the swelling percentage was measured at each time point, and gel strength was evaluated along with the degradation.

[0072] In vivo durability testing Isolation of canine pancreatic islets Canine islets were isolated from locally obtained pancreata from euthanized donors at a local veterinary hospital with owner consent for organ donation. Procurement and digestion protocols have been previously published in detail. Organs were harvested and washed, followed by collagenase digestion followed by density gradient purification using standard published protocols. Isolated islets were converted to islet equivalents, or "IEQ", for quantification purposes by dithizone staining following standard published protocols. Canine islets were cultured at 37°C and 5% CO2 in CMRL1066 supplemented with 10% fetal bovine serum, 2 mM glutamine, 10 mM nicotinamide, and 1% antibiotic-antimycotic solution.

[0073] Xenotransplantation of Encapsulated Pancreatic Islets Immunodeficient NOD / SCID mice (NOD.CB17-Prkdc) were treated with streptozotocin (STZ, 220-250 mg / kg) to induce diabetes. scidThe functional capacity of CSS-encapsulated canine islets was evaluated using a CSS-encapsulated islet cytochrome P450 (CSS-C) ELISA Kit (Jackson Laboratory). Six mice received islets encapsulated in PEGDA, two mice received islets in PEGMAL (synthetic procedure below), and four mice received the same dose of islets encapsulated in MeHA. Two mice were transplanted (intraperitoneally) with an equivalent dose of non-encapsulated islets as controls. At week 10 of the study, all mice receiving islets encapsulated in MeHA became hyperglycemic and were terminated along with the non-encapsulated islet group. Four mice from the PEGDA group were also terminated at week 10 to provide a comparison for later histological studies. The remaining two PEGDA mice were terminated after an additional six weeks. The two PEGMAL mice are still being monitored (>16 weeks after transplantation).

[0074] Durable PEGDA and fast-degrading MeHA formulations of islet-encapsulated microparticles were prepared as described above, except that canine islet cells were mixed into the precursor. Chemically crosslinked (i.e., non-UV crosslinked) durable PEGMAL formulations were prepared following a similar scheme to ThHA microparticles described above. Briefly, hydrogel precursors consisting of 25% (w / w) 8-branched 40 kDa PEGMAL (JenKemUSA) and canine islets were extruded into a pH 6.6 alginate bath containing 1 kDa SH-PEG-SH crosslinker (BiochemPEG) at a concentration of 0.25% (m / v) and stirred for 5 min. After 5 min, the crosslinking alginate bath was diluted in half with HEPES-buffered islet medium to a final pH of approximately 7.2 and stirred for an additional 30 min to allow complete crosslinking of the hydrogel core. The alginate shell was then removed as described.

[0075] Islet-containing microparticles were administered intraperitoneally (IP) to mice via an 18G catheter. The islet dose was based on a previous ascending dose study showing that approximately 4000 islet equivalents / mouse resulted in normoglycemia and is consistent with previous publications in which canine islets were transplanted into mice. Blood glucose levels were measured daily for 14 days after transplantation and then biweekly for all mice.

[0076] After the animals were sacrificed (10 and 16 weeks), islet-containing microparticles were collected from the peritoneal cavity and plated in culture medium for evaluation. The islet-containing microparticles were stained for the presence of insulin by dithizone staining, and the cell viability of the explants was evaluated by calcein staining (viable cells) and propidium iodide staining (apoptotic / necrotic cells). Color, bright-field, and fluorescent images of the stained microparticles were taken using a Cytation5 Imaging Multimode Reader (Biotech Instruments, Inc.).

[0077] Tissue samples from the pancreas and particulate-bearing peritoneum were harvested and fixed overnight in 10% neutral buffered formalin, then transferred to PBS. Tissues were embedded in paraffin blocks and appropriately sectioned for staining. For hematoxylin / eosin (H&E) staining, sections were sequentially deparaffinized by placing them in Clear Rite3 solution three times for 3 min each, followed by dehydration and rehydration steps before staining with hematoxylin and eosin. Sections were rinsed with water, followed by addition of Bluing Reagent, and further rinsed with tap water. Finally, sections were immersed in 100% absolute alcohol and Clear Rite 3 to improve color stability, and coverslipped as appropriate. Slides were examined and images were captured with a Biotek Cytation5 cell imaging multimode reader.

[0078] Allogeneic transplantation of encapsulated islets Two adult male dogs recently diagnosed with diabetes received canine islet grafts encapsulated with either durable PEDGA or rapidly degrading MeHA (N=1 / group). Canine islet encapsulation followed a previously described protocol. Each dog received a graft consisting of approximately 60 mL of preloaded microparticles delivered to the peritoneum via direct catheter injection. The dogs were monitored daily for weight and blood glucose for the first 20 days after transplantation.

[0079] result In vitro degradation MeHA, PEGDA, and AHA microparticles were placed in HEPES-buffered degradation solution at 37°C for 1, 3, or 6 weeks, then dried and the dry polymer mass was determined. After 6 weeks of degradation, MeHA microparticles were found to retain only 24% of their starting mass, while PEGDA and AHA microparticles retained 37% and 53%, respectively (Figure 8A). Although PEGDA lost a significant amount of mass (Figure 8C), it maintained its gel strength over the 6-week period, as evidenced by the unchanged swelling ratio (Figure 8B). In contrast, MeHA and AHA microparticles showed a dramatic increase in swelling ratio, indicating a significant decrease in gel strength as the microparticles degraded. The difference in gel strength between PEGDA and HA-based microparticles can be attributed to both crosslink density and polymer chemistry. In the case of PEGDA microparticles, the polymer chains are tightly held together by several covalent bonds at a higher crosslink density compared to HA microparticles. In addition, the PEGDA polymer chains contain several regions that are hydrophobic moieties. Both the crosslink density and hydrophobic regions allow water to diffuse into the microparticles at a slower rate than hydrolysis occurring on the surface of the microparticle. Of course, if degradation were only on the surface, the internal gel network would remain intact and gel strength would be preserved. In MeHA and AHA microparticles, however, the crosslink density is much lower compared to PEGDA, and the polymer chemistry results in a more hydrophilic structure. The lower crosslink density and hydrophilicity of HA microparticles both favor a higher rate of water diffusion into the microparticles, resulting in a bulk degradation mechanism. In bulk degradation, water penetrates deeper into the hydrogel network, resulting in hydrolysis that compromises the strength of the gel network. Such degradation may explain why AHA microparticles retain a higher polymer mass than PEGDA after 6 weeks of degradation. The decrease in gel strength over time in HA microparticles indicates that hydrolysis is occurring, but the degradation products have not yet diffused out of the gel network. In this case, the remaining polymer mass would not be an accurate assessment of the extent of degradation, since the polymer is still present inside the gel network but no longer contributes to the integrity of the hydrogel.

[0080] Function of canine islet-loaded microparticles in diabetic mice Canine islets encapsulated with PEGDA or MeHA were transplanted into diabetic NOD / SCID mice. Control groups received nonencapsulated islets. Figure 9 shows the average daily blood glucose levels for each group in the initial study cohort after transplantation. Mice in the MeHA, PEGMAL, and PEGDA groups were all normoglycemic by the second week after transplantation. However, animals receiving the rapidly degrading MeHA microparticles gradually reverted to hyperglycemic levels (beginning approximately 3 weeks after transplantation) despite the reintroduction of exogenous insulin treatment. These mice were euthanized around the 10-11th week. Mice receiving durable PEGDA and PEGMAL encapsulated islets remained normoglycemic throughout the entire study period (PEGMAL ongoing, 16 weeks), providing excellent glycemic control without exogenous insulin.

[0081] In contrast, none of the mice receiving nonencapsulated islets achieved sustained normoglycemia at any time point. These mice had an initial mean blood glucose of 386 ± 23 mg / dL before transplantation and 494 ± 61 mg / dL on the day of transplantation. After transplantation of the same volume of islets as the other two groups, blood glucose showed only one normal measurement on day 2 in one animal, while the other mice showed fluctuations in blood glucose from days 19 to 30, but then returned to consistently hyperglycemic levels for the remainder of the study.

[0082] Microparticles were recovered from the abdominal cavity of the mice at necropsy. The results are shown in Figures 10A-10D. The durable PEGDA microparticles containing islets were easily recovered around week 10, whereas the rapidly degrading MeHA microparticles were very few in number and difficult to identify. Recovered samples of PEGDA microparticles were found to be irregularly distributed throughout the abdominal cavity and remained transparent and structurally intact. Recovered PEDGA microparticles showed little to no signs of degradation (i.e., smooth edges, no obvious change in size, mechanically stable) and the transplanted cells were still contained within the microparticles (Figure 10A). In contrast, the rapidly degrading MeHA microparticles were misshapen and small in diameter, indicative of significant hydrogel degradation. The MeHA microparticles did not appear to contain intact islets (Figure 10B). At later time points (week 16), the durable PEDGA microparticles continued to be easily recovered at necropsy. The islets within the PEGDA microparticles were stained with dithizone, calcein, and propidium iodide (PI) to confirm the presence of viable and functional islets.

[0083] PEGDA microparticles harvested around 16 weeks post-implantation contained healthy, live islets, identified by deep red dithizone staining (Figure 10C). The same islets were co-stained with green calcein, a common viable cell indicator (Figure 10D). Some positive PI staining (red; apoptotic / necrotic cells) was observed, but was generally localized to non-islet (i.e., dithizone-negative) tissue that appeared attached to the surface of the harvested microparticles. Conversely, islets (dithizone-positive cells) were almost completely free of PI (dead cell) staining (Figure 10D).

[0084] The durable PEGDA microparticles were found to adhere to tissues such as the liver and abdominal wall, but were generally nonadherent and floated freely within the cavity. Histological examination of the graft sites identified the location of the PEGDA microparticles. Although the process for processing for histological examination destroyed the integrity of the PEGDA microparticles, there was no obvious fibrotic tissue at the attachment site. Figure 11 shows a healthy adipocyte crossed by two microparticles, whose locations are indicated by dark lines. The thin bluish-grey layer in the center of each microparticle location is the hydrogel that remained after processing. In contrast, no gel material could be identified in the histological sections obtained from the MeHA group. In the PEGDA group, generally no fibrosis was seen at the interface of the microparticles and the retina. To confirm that the normalization of blood glucose levels in PEGDA-treated mice was due to the engraftment of the graft and not due to endogenous islets remaining in the pancreas after streptozotocin treatment, histological sections of the pancreas were examined for the presence of islets. Only one islet-like structure was identified in six pancreatic sections per mouse (results not shown).

[0085] A similar study was completed with canine islets transplanted into two diabetic dogs. Both animals were diabetic and did not receive regular exogenous insulin prior to transplantation. One dog received canine islets encapsulated in durable PEGDA microparticles, while the other received canine islets encapsulated in degradable MeHA hydrogel. Figure 12 shows the mean non-fasting blood glucose from measurements taken each morning and afternoon. The dog receiving islets in durable hydrogel slowly returned to lower blood glucose levels within the normal range (solid line) and maintained those normal levels for the duration of the experiment (8 weeks). In contrast, the animal receiving the rapidly degrading hydrogel slowly returned to a fully diabetic state within 20 days of transplantation (dotted line).

[0086] Consideration Interestingly, both groups of mice achieved normoglycemia after intraperitoneal injection of the microparticles, whereas nonencapsulated islets at the same dose and implantation site did not. Given that the animal model used immunodeficient mice, this result suggests that the hydrogel matrix provides a more physically supportive environment that promotes better cell function and survival, especially in the context of implantation via intraperitoneal injection. This conclusion is supported by the observed degradation of MeHA microparticles over time, which coincided with the return of hyperglycemia. More notably, PEGDA and PEGMAL islet microparticles were structurally more robust and therefore provided better glycemic control compared to MeHA transplantation during the peri- and post-transplant periods before hyperglycemia recurred in the MeHA group.

[0087] Although only two animals were used in the dog study, the results are important because they contrast the differences in responses between immunocompromised mice and immune-competent dogs. The intact dog immune system caused the more fragile hydrogel (MeHA) to degrade more rapidly, resulting in a faster return to hyperglycemic blood glucose levels. The results indicate that this particular degradable hydrogel microparticle formulation has an in vivo persistence of approximately days to weeks, depending on the species. However, the durable hydrogel microparticles demonstrated an in vivo persistence of at least two months.

[0088] Example 3 Comparison of in-bead growth of durable and rapidly degradable particulate end-products. Introduction Manipulation of the physical and chemical properties of cell encapsulation materials has become a powerful tool for in vitro tissue engineering research. The mechanical and chemical properties of the microenvironment are known to alter the behavior of various cell types, most notably stem cells, which are influenced by the mechanical properties of their surroundings. The ability of stem cells to continue to proliferate within hydrogels is an important property in the therapeutic treatment of soft tissue defects due to trauma or surgery. For example, muscle or skin injuries would benefit from the local delivery of microbeads containing proliferating stem cells. Proliferating stem cells allow for a continued expansion of cell numbers while present within the microparticles, thus allowing for smaller microparticle dosages. However, not all hydrogels allow for intramicroparticle proliferation of cells. Here, we show that rapidly degrading hydrogel microparticles enable robust cell proliferation within the microparticles.

[0089] method Human embryonic kidney cells (HEK293) were cultured in DMEM medium containing high concentrations of glucose and fetal bovine serum and maintained in an incubator at 37 °C and 5% CO2. Once the cells reached an appropriate density, they were detached from the culture flask by adding 0.5% trypsin-EDTA for 2 min. Immediately after detachment, the cells were transferred to growth medium. To test the ability of cells to grow within the hydrogels, microparticles were fabricated from two hydrogel polymers, the fast-degrading MeHA and the durable PEGDA, using the CSS fabrication method described above. The same amount of polymer precursor (1 mL) was added to the same number of proliferating human embryonic kidney cells (HEK293; 4 × 10 6 The cells were mixed with the cellulose membrane and formed into microparticles using the method described above. The microparticles were maintained under culture conditions for 3 weeks. To evaluate the proliferation of cells within the microparticles, images were taken over a 34-day period using a Biotec Cytation5 cell imaging multimode reader.

[0090] result Upon incorporation of microparticles, we found that cell morphology and intra-microparticle distribution were unique depending on the hydrogel. Initially, in durable PEGDA microparticles, HEK293 cells were clustered (Figure 13A). In contrast, the same cells loaded at the same density in degradable (MeHA) microparticles remained as single cells (Figure 13C).

[0091] Over time, there was little change in cell morphology or number within the PEGDA microparticles (Figure 13B). In contrast, cells within the degradable MeHA microparticles proliferated within the hydrogel and formed large clusters within the microparticles that were visible after 34 days in culture (Figure 13D). These clusters did not appear to protrude from or attach to the surface of the MeHA beads, but remained encapsulated within the bead matrix. However, the same cells within the durable PEGDA microparticles did not proliferate rapidly or form proliferative cell clusters within the PEGDA beads.

[0092] Consideration In these final product forms, each microparticle either prevents or allows cell proliferation within the microparticle. As shown by the data presented in this example, the durable hydrogel prevented cell proliferation, while the softer, rapidly degrading MeHA hydrogel allowed significant proliferation of fetal cells. In addition, cell morphology changed depending on the formulation used. This may be important when encapsulating proliferative versus non-proliferative cells, and can be tuned using CSS technology.

[0093] Example 4 Comparison of cell migration on durable and degradable particulate end products Introduction In several therapeutic applications, extracellular migration of cells and their products is an important aspect of treatment. For example, endogenous stem and progenitor cells have been shown to migrate to the brain during normal development and also under pathological conditions such as brain injury and disease. Some studies have suggested that native stem cells may actually migrate or "hone" into the injured area to aid in its repair. However, most studies have been unable to substantiate such claims when controlled by in situ measurements of stem cell numbers in uninjured tissue.

[0094] That is, for cells to help repair any tissue, they need to be localized in that area for an extended period of time. The therapeutic practice of placing unencapsulated stem cells in an area is challenging because the cells are small and can quickly diffuse out of the area or be attacked by the immune system. Thus, a localization approach that allows for the sustained release of cells to a specific area would be of great therapeutic benefit. In this example, we demonstrate these properties in degradable microparticles.

[0095] method Three hydrogel polymers, MeHA, AHA, and PEGDA, were fabricated into microparticles using the CSS fabrication method described above. Human embryonic kidney cells (HEK293) were cultured in DMEM medium containing high concentrations of glucose and fetal bovine serum and maintained in an incubator at 37 °C and 5% CO2. Rat bone marrow stem cells (rBMSCs) were cultured in high glucose DMEM, fetal bovine serum, and Glutagro. Once the cells reached an appropriate density, they were detached from the culture flask by adding 0.5% trypsin-EDTA for 2 min. Immediately after detachment, the cells were transferred to growth medium. The same amount of polymer precursor (1 mL) was added to the same number of HEK293 cells or rBMSCs (4 × 10 6 ) and then bead formation was carried out.

[0096] To assess the amount of cell migration out of the microparticles, at the indicated time points, the microparticles were removed from the medium and washed across a screen. The wash medium and the medium surrounding the microparticles were collected. Cells were counted using both a Cytation5 Imaging Multimode Reader (BioTek Instruments) and an EVE Cell Counter.

[0097] result Figure 14 shows typical images of wells containing collected spent and wash media containing individual cells 4 days after encapsulation. Few cells were counted in the media of HEK293 cells encapsulated in PEGDA hydrogels, many of which were dead (red) (Figure 14A). In contrast, the degradable hydrogels allowed the release of a large number of live cells, as shown by the large number of live cells (green) remaining in the wells in Figure 14B. However, these cell numbers were small compared to the accumulated cell release values ​​at 2 weeks.

[0098] Figure 15 summarizes the average data from the cell release accumulation study. During the first 6 days of culture, only a few thousand cells were released from the beads. By day 14, 2 x 10 7 More than 10 cells were released by the rapidly degradable MeHA microparticles, whereas only 2.2 × 10 cells were released from the durable PEGDA beads. 5 Cumulatively, by day 21, MeHA beads released 3.08 × 10 cells. 7 cells were released, whereas PEGDA released less than 5% of the cells.

[0099] Consideration The results show a clear difference in the number of proliferative cells migrating out of the durable microparticles compared to the number of proliferative cells migrating out of the degradable microparticles. The final microparticle product allows cells to migrate out of the microparticles and into the surrounding tissue. The rapidly degrading formulation allows for faster and greater migration than the durable hydrogel. These results demonstrate the utility of rapidly degrading hydrogels in allowing cells to migrate to the local injury site even before physical degradation of the microparticles has occurred.

[0100] Example 5 Comparison of viability, morphology, and functionality of durable and rapidly degradable particulate end products Introduction There are numerous applications for durable and fast-degrading / short-term hydrogel formulations. Literature has shown that microencapsulated mesenchymal stem cells (MSCs) dramatically enhance paracrine-mediated healing in a mouse hind limb injury model, and demonstrate increased viability and proangiogenic activity when compared to nonencapsulated MSCs. Once encapsulated, the cells consistently maintain viability and are consistently functional therapeutically. Increasing the stiffness of the encapsulation matrix has been shown to drive MSC differentiation toward a more osteogenic pathway, whereas more flexible materials drive the differentiation toward a chondrogenic pathway. Furthermore, rat neural cells cultured in three-dimensional collagen hydrogels exhibited better viability and behaved more similarly to native neural networks compared to cells grown on the same materials in two dimensions.

[0101] method Viability and morphology Two types of hydrogel polymers, one rapidly degradable (MeHA and AHA) and one durable (PEGDA), were fabricated into microparticles using the CSS fabrication method described above. The same amount of polymer precursor (1 mL) was mixed with the same number of mouse MSCs (1.6 × 10 7 cells) or insulinoma cells (3.0 × 10 7 The microspheres were mixed with 1000x1000 cells (100x1000 cells) and then bead formation was performed. The beads were maintained under culture conditions for 2 weeks. The viability and functionality of the cells when cultured in the microspheres was high for both formulations. After encapsulation, the viability of MSCs and insulinoma cells in the microspheres was measured using a fluorescent viability stain. Calcein was added to the medium as a measure of live cells, since it can only enter cells with an intact membrane. To identify dead cells, propidium iodide was added to the medium. After 30–60 min of incubation in the fluorophore, images of the microspheres were collected with a Cytation5 imaging multimode reader (Biotech Instruments).

[0102] Similar experiments were also performed using a non-proliferating somatic cell source (canine islets) encapsulated in two different hydrogels. Microparticles with cells (islets or cultured cells) mixed into the precursor were produced as described above. For canine islets, a 1.1x concentration of precursor was mixed with the canine islet slurry in a 10:1 volume ratio immediately followed by droplet generation. The production of islet microparticles was performed aseptically in a closed, sterile bioreactor system.

[0103] The completed microparticles were stained for the presence of insulin by dithizone staining, and further assessed for cell viability in the explants by calcein staining (viable cells) and propidium iodide staining (apoptotic / necrotic cells). Color, brightfield, and fluorescent images of the stained microparticles were captured using a Cytation5 Imaging Multimode Reader (Bio-Tek Instruments, Inc.).

[0104] result As shown in Figure 16A and Figure 16B, MSCs showed high viability when encapsulated in either PEGDA or MeHA. Similar results were seen with AHA (Figure 16E). Green fluorophores indicate live cells stained with calcein. Red / yellow cells indicate dead cells stained with propidium iodide. Interestingly, cells were loaded into the microparticles as single cells, but while they rapidly clustered in the durable PEGDA gel, no clustering occurred in the degradable MeHA hydrogel and they remained mainly as single cells. Monitoring of cell morphology during the fabrication process indicated that the cells clustered during microparticle fabrication, not after fabrication.

[0105] In studies with canine islets, cells had high viability in both hydrogel microparticles (Figure 16C: PEDGA, Figure 16D: MeHA). Cells remained functional in both hydrogels as shown by cell staining for insulin content. Islets were loaded within the microparticles as clusters of cells. There appeared to be no change in clustering frequency or cluster size in either formulation.

[0106] Similar results were obtained with a rat insulinoma cell line. Figure 17 shows that cell viability is high, especially when encapsulated in AHA, within the first day of encapsulation. Over the next 7 days, cell viability in AHA dropped to approximately 50%. In contrast, insulinoma cells in PEGDA started out with a lower viability but remained near the mean throughout the study.

[0107] The functionality of the islet cells was tested by staining for intracellular insulin. Dithizone staining is a standard indicator of insulin production. Dithizone staining stains insulin positive cells with a red / brownish hue. The images in Figure 18 show that the islets stain positive for insulin approximately one week after encapsulation in both durable PEGDA and degradable MeHA microparticles, indicating that insulin production can continue after encapsulation in either microparticle. Unlike mouse MSCs, the morphology of these somatic islet cells remained largely unchanged in either hydrogel formulation.

[0108] Example 6 Comparison of local cellular administration of durable versus rapidly degradable microparticle end products. Introduction Durable and rapidly degradable hydrogel formulations have numerous applications. It has been shown in the literature that microencapsulated MSCs dramatically enhance paracrine-mediated healing in a mouse hind limb injury model, demonstrating increased viability and proangiogenic activity when compared to non-encapsulated MSCs. One issue with the use of paracrine release of MSCs is that unencapsulated MSCs do not reside in the area of ​​injury, but are rapidly removed from the body. Thus, encapsulation with an adhesive hydrogel allows the cells to be trapped in the area of ​​implantation where they can secrete beneficial compounds. Experiments were performed to determine whether durable or degradable microparticles exhibited a difference in their ability to attach to the surrounding tissue and remain in the area.

[0109] method Adhesion to tissue Immediate tissue adhesion was examined during transplantation into the canine retina. Four dogs were transplanted with microparticles containing canine islets. On the day of transplantation, the animals were anesthetized using standard veterinary procedures, including a cephalic vein catheter for fluid injection before, during, and after the surgical procedure. The microparticles were injected into the retina by the surgeon. Care was taken not to penetrate or damage any of the intestinal tissue, blood vessels, lymph nodes, or lymphatics. A total of 10,000 islet equivalents were contained within 1 mL of hydrogel microparticles. The preparation of islet-containing PEGDA microparticles was previously described. After photographs were taken, the retina was gently placed back into the abdomen, the abdominal wall was closed, and then the skin was closed with intradermal sutures.

[0110] As another example, tissue adhesion was demonstrated by injecting microbeads into rat knees. Briefly, 100 μL of 300 μm PEGDA microparticles in PBS were extruded through a 25G needle into the joint capsule of a Sprague-Dawley rat knee joint. To confirm the presence of the microparticles inside the joint capsule, the solution and beads were stained blue prior to extrusion.

[0111] Histological examination of adherent tissue Healthy Sprague-Dawley rats were used to assess acute biocompatibility and intraperitoneal microparticle location (N=2 per microparticle formulation). Empty microparticles were implanted around the retina as previously described. Microparticles were delivered by syringe as a suspension in DPBS. Approximately 1 mL of loosely packed microparticles were allowed to accumulate around and within the rat retina. Rats were monitored and scored daily for pain markers and activity levels for 10 days after microparticle implantation.

[0112] Fourteen days after implantation, rats were euthanized and the implantation sites were evaluated to determine the location of microparticles in the abdominal cavity and to examine for possible tissue abnormalities. Tissue samples were collected, stored in neutral buffered formalin, embedded in paraffin, and sectioned at 7 μm thickness. Sections were stained with H&E and evaluated microscopically.

[0113] result As shown in Figure 19A and described in the Methods section, PEDGA microparticles were injected into and around the ventral retina. Immediately after injecting the microparticles in vehicle into and around the retina, images were collected showing that the microparticles adhere to the tissue even when the retina is manipulated. Figures 19B and 19C show results from two separate dogs, both of which demonstrate the adhesive nature of the microparticles.

[0114] Dyed PEGDA microparticles were injected into the knees of rats. The joints were manipulated and then dissected. The blue color from the microparticles was visible through the muscle (Figure 20A). Upon further dissection, the microparticles were visible and still present within the knee joint (Figures 20B and 20C).

[0115] Durable PEGDA microparticles, degradable MeHA microparticles, and degradable AHA microparticles (without cells) were implanted into healthy Sprague-Dawley rats to assess the safety and biocompatibility of microparticles produced using the CSS method, as well as their location and adhesion to surrounding tissues. Necropsies performed at 2 and 10 weeks revealed no signs of acute inflammation, excess fluid, or tissue abnormalities anywhere in the abdominal cavity, although some tissue adhesions were noted where the retina had been sutured to the stomach wall. Both microparticle formulations were found intact in the retina, were visually translucent, and appeared to be surrounded by a very thin membrane-like layer. A few microparticles were found attached to the surface of the liver. However, the majority of the microparticles were attached to the retina, with a minority being free-floating.

[0116] Hematoxylin and eosin staining of explant tissues for the three microparticle groups is shown in Figure 21. No significant differences were observed between the groups. Microparticles were generally intact, but appeared to be shrunken in size and in some cases collapsed / distorted as a result of tissue processing for histology. Microparticles were commonly found to have a thin cell layer around their periphery, often 2 or 3 cells thick, and all microparticles were devoid of cells as empty microparticles were injected into the animals. Most single microparticles were surrounded by normal retinal tissue, indicating a lack of inflammatory response in response to the various formulation microparticles.

[0117] Quantification of the foreign body response of the microparticles in the retina is shown in Figure 22. The width of the fibrotic region, defined by low cellularity and high collagen content, was minimal for AHA and elevated around PEGDA microparticles. Nevertheless, the thickness of the fibrotic ring around the PEGDA microparticles was less than that reported for other hydrogel formulations such as alginate. Similarly, the lymphocytic region was smaller for AHA microparticles. The lymphocytic region was defined as the area where cellular immune cells had migrated around the microparticles. The lymphocytic region was also higher for PEGDA microparticles, but was still considered minimal (Figure 22).

[0118] Consideration A unique beneficial property of the final product is that it is adhesive, whether composed of a durable or degradable hydrogel matrix, which allows the final product to rapidly attach to surrounding tissue and remain in the area until degraded. In addition to the attachment of islets to the surrounding tissue in the peritoneal cavity, the adhesive hydrogel was used to administer cells such as MSCs that release anti-inflammatory and pro-healing cytokines, allowing for long-term delivery of cellular factors to a local area via injection into inflamed tissues such as joints, without concern for redistribution to other areas of the body.

[0119] Example 7 Examination of parameters affecting hydrogel properties In this example, various parameters were explored to better understand their relationship to one another and their effect on the properties and characteristics of the resulting hydrogel microparticles. As discussed in more detail below, various microparticle characteristics can be finely tailored to specific desired treatment protocols and outcomes. Notably, each of these parameters is a dynamic factor in hydrogel microparticle production, meaning that its effect is related to the amounts or ratios of other components used in the formulation and / or other processing parameters. Furthermore, changes to a particular parameter do not necessarily result in a proportional change in the resulting hydrogel. Nevertheless, by nuanced variations in the various microparticle formulations and processing parameters, many different characteristics can be customized depending on the desired outcome, such as the type of cells to be encapsulated, the location of implantation, and the condition to be treated or prevented.

[0120] To investigate these various parameters, the swelling ratio (Q) is used as a surrogate for the relative degradation rate of the microparticles.

[0121] A. Mass fraction of core polymer species Mass fraction refers to the mass % composition of the reactive polymer species in the hydrogel precursor solution used in the polymer backbone (e.g., for 2.5 g PEGMAL in 10 g precursor solution, the mass fraction is 25%). Adjusting the mass fraction of the reactive polymer in the precursor solution has a consistent effect on the resulting Q value. The lower the mass fraction, the higher the Q value and the softer the gel. This is demonstrated by crosslinking various hyperbranched PEGMAL species with a 500 Da PEG dithiol. The crosslinker concentration in this example was held constant at 0.25% mass fraction in the alginate bath. A non-reactive PEG species (PEG8000) was included in the low mass fraction group to maintain a constant solution viscosity. The results are shown in Figure 23.

[0122] B. Concentration of cross-linking species in the alginate bath The concentration of crosslinker (e.g., PEG dithiol, DTT) in the alginate bath influences the resulting hydrogel microparticles. However, the relationship between crosslinker and hydrogel matrix is ​​not simple, as there can be too little or too much crosslinker in the bath. Insufficient crosslinker will result in a poorly crosslinked, brittle gel; too much crosslinker will saturate the reactive sites on the polymer backbone, similarly preventing crosslinking and resulting in a brittle gel. That is, this parameter must also be balanced with the amount and type of polymer precursor used in the backbone.

[0123] 1. Increasing the concentration of crosslinker decreases the Q value. In this example (core polymer = 4-branched 10 kDa PEGMAL), lower Q values ​​were obtained when the crosslinker concentration (DTT) was increased, indicating that the crosslinker concentration never reached the level at which saturation of the core polymer occurs. The results are shown in Figure 24A.

[0124] 2. Increasing the crosslinker concentration increases the Q value. In this example (core polymer = 8-branched 40 kDa PEG-vinylsulfone), increasing the crosslinker (BMA, CAS123-81-9) concentration initially lowered the Q value, but further increase led to higher Q values, in contrast to what was observed in Example 1. Of note, the relative molar concentration of crosslinker (thiol) to core reactive group (vinylsulfone) was different from the previous examples. That is, the conditions in Example 2 may have led to saturation of the core polymer molecules, leading to insufficient crosslinking, resulting in a weaker gel network and higher Q. The results are shown in Figure 24B.

[0125] C. pH of the alginate bath The thiol-ene reaction is strongly influenced by the pH of the environment: increasing the pH value dramatically increases the thiol-ene reaction kinetics, i.e., changing the pH of the alginate bath can have a significant effect on the crosslinking dynamics of the gel and the resulting Q value.

[0126] 1. Increasing the pH increases the Q value. In this example (core polymer = 8-branched 40 kDa PEG-vinylsulfone; crosslinker = BMA), increasing the bath pH resulted in higher Q values. The results are shown in Figure 25A. The increased reaction rate may have led to a faster reaction and ultimately to partial saturation of the core polymer, resulting in a weaker gel network and higher Q values.

[0127] 2. Increasing the pH decreases the Q value. In this example (core polymer = 8-branched 40 kDa PEGMAL; crosslinker = 500 Da PEG dithiol), increasing the pH resulted in lower Q values. The polymer and crosslinker are balanced to avoid saturation; that is, increasing the pH in this case resulted in faster reaction kinetics and more efficient crosslinking. The results are shown in Figure 25B. Note that the increase in Q value with increasing pH is much smaller in Example 2 compared to the very sharp increase in Example 1. That is, both the direction and magnitude of the effect of pH on Q value appear to be strongly influenced by the relative amounts of crosslinker and core polymer reactive sites.

[0128] D. Molecular weight of crosslinking species The molecular weight of the crosslinking species affects both the rate of diffusion into the core-shell construct and the distance between crosslinks within the gel network, i.e., differences in the molecular weight of the crosslinking species can affect the final Q value of the resulting gel particulates depending on other parameters used during the process.

[0129] 1. Increasing the molecular weight increases the Q value. In this example (core polymer = 8-branched 40 kDa PEG-vinylsulfone), increasing the molecular weight of the crosslinkable species from 154 Da to 500 Da significantly increases the Q value. The molar concentration of the crosslinker species was 10 mM for both preparations. The results are shown in Figure 26A.

[0130] 2. The Q value does not change even if the molecular weight is increased. In this example (core polymer = 8-branched 40 kDa PEGMAL), there was no obvious difference in the Q value of the resulting gel when the molecular weight of the crosslinker was increased. The effect of changing the molecular weight of the crosslinker may offset the adjustment of the relative amount of crosslinker used to the polymer reactive sites. The results are shown in Figure 26B.

[0131] E. Molecular Weight of Core Polymer Species Assuming the number of reactive sites per gram of polymer is conserved, the molecular weight of the core polymer species appears to have a much more predictable effect on Q than the molecular weight of the crosslinker. For example, an 8-arm 40 kDa PEGMAL and a 4-arm 20 kDa PEGMAL have the same number of reactive sites (maleimides) per gram. However, as shown below, the 40 kDa species yields much stronger gels (lower Q) when prepared under otherwise identical circumstances.

[0132] 1. Effect of molecular weight of core polymer at constant number of reactive sites / gram In this example, three side-by-side comparisons are used to demonstrate the effect of core polymer molecular weight on the Q value of gel microparticles produced by CSS. In all preparations, increasing core molecular weight decreased the Q value even at the same mass fraction and number of reactive sites per gram. Note that the effect of core molecular weight becomes more pronounced with decreasing crosslinker concentration (and increasing size). The results are shown in Figure 27A.

[0133] 2. Effect of molecular weight of core polymer when number of reactive sites / gram is not equal In this example, microparticles consisting of 8-branched 40 kDa or 4-branched 10 kDa PEGMAL core species are compared. In all cases, the core polymer solution was at 10% mass fraction. Although the 4-branched 10 kDa species has twice the number of reactive sites / gram when compared to the 8-branched 40 kDa, the 8-branched 40 kDa microparticles had lower Q values ​​(i.e., more rigid gels) compared to the 4-branched 10 kDa microparticles, despite the lower potential crosslinking density. The results are shown in Figure 27B.

[0134] F. Effect of UV curing time In this example, AHA and MeHA microspheres of various molecular weights were produced with various UV exposure times. Briefly, 2.5% solutions of 1 MDa or 200 kDa AHA or 1 MDa MeHA (each containing 1% 3.4 kDa PEGDA) were added dropwise by extrusion to a stirred alginate bath as described above and exposed to UV light for 2.5, 5, or 7.5 minutes. The resulting diameter and hydrogel swelling ratio for each polymer type were quantified. The results are shown in Figures 28A and 28B, which show the change in microsphere diameter and swelling ratio as a function of UV exposure time. It is clear from these results that as UV exposure time increases, both the diameter and swelling ratio decrease. Notably, the swelling ratio of 1 MDa MeHA hydrogel microspheres is dramatically reduced after 7.5 minutes of UV exposure compared to 5 minutes, with swelling ratios of 509.1 and 113.4, respectively. This trend was consistent across polymer types and molecular weights, demonstrating control over hydrogel durability by varying UV exposure time.

[0135] The hydrolytic stability of the crosslinking species may control hydrogel degradation, independent of the GQ value. 1. BMA as a more easily hydrolyzable crosslinker. Certain crosslinkers can be used to affect the degradation properties of microparticles. For example, the dithiolate crosslinker molecule BMA could be used as a means to enhance the hydrolytic degradation of thiol-ene hydrogels.

[0136] A series of hydrogel microparticles composed of PEG-vinylsulfone crosslinked with DTT or BMA were prepared and stored in DPBS at room temperature (approximately 27°C) in 50 mL polypropylene tubes for at least 6 months. Table 3 below lists the composition and corresponding swelling ratio of each preparation for reference. After 6 months of storage, each microparticle was qualitatively re-examined for signs of degradation. Of note, all microparticles crosslinked with BMA were completely degraded by 6 months. Those crosslinked with DTT appeared relatively unchanged after 6 months, but formal characterization was not performed at this time. Furthermore, two of the fully degraded BMA crosslinked microparticles previously exhibited the lowest Q values ​​of all microparticles included in this table, but still showed complete degradation when compared to the higher Q value DTT crosslinked microparticles. This result suggests that the BMA crosslinker may be a useful tool to generate microparticles that are initially very robust (low Q) but degrade much more rapidly than other microparticles of similar initial strength crosslinked with DTT or PEG dithiol. [Table 3]

[0137] Table 4 below lists additional dithiol species and the corresponding pKa of the thiol group. The hydrolytic lability of the resulting bond in the thiol-ene reaction correlates with the pKa of the reactive thiol. Thus, these species can be used as chemical crosslinkers in thiol-ene reaction schemes to modulate the hydrolytic lability of the microparticles and therefore their degradation rate. [Table 4]

Claims

1. 1. A method for the localized delivery and sustained release of therapeutic cells and / or tissues at a site of implantation comprising the steps of: forming a non-alginate based hydrogel microparticle having a diameter greater than about 100 mm; forming a non-alginate based hydrogel microparticle having a diameter greater than about 100 mm; and forming a non-alginate based hydrogel microparticle having a diameter greater than about 100 mm.

2. The method for the localized delivery and sustained release of therapeutic cells and / or tissues at a site of implantation comprising the steps of: forming a non-alginate based hydrogel microparticle having a diameter greater than about 100 mm;

2. 10. The microparticle of claim 1 having a size greater than 300 μm.

3. 10. The microparticle of claim 1, wherein the matrix consists essentially of covalently crosslinked polyethylene glycol maleimide and polyethylene glycol dithiol precursor compounds.

4. The microparticle of claim 1 , comprising up to 50% of the cell or tissue by volume of the microparticle.

5. 2. The microparticle of claim 1, having a storage stability in PBS at 37°C for less than 6 months.

6. 2. The microparticle of claim 1, having a storage stability in PBS at 37°C for more than 6 months.

7. The microparticle according to claim 1, which is a degradable microparticle that decomposes in less than three months when implanted in a living body.

8. The microparticle according to claim 1, which is a durable microparticle that does not decompose for at least three months when implanted in a living body.

9. The microparticle of claim 1 further comprising a cell culture medium component.

10. The microparticle of claim 1 , wherein the cells are non-proliferating cells or proliferating cells.

11. 10. The microparticle of claim 1, further comprising, in addition to cells, secreted cell products, such as signaling molecules, cytokines, chemokines, therapeutic proteins, hormones, vesicles, antibodies, viruses, exosomes, and the like.

12. The microparticle of claim 1 , wherein the cell is a genetically engineered or transgenic cell.

13. The microparticle according to claim 1 , wherein the cells are a cell mass consisting of a plurality of aggregated cells.

14. The microparticle of claim 13 , wherein the cell mass comprises two or more different types of cells.

15. 10. The microparticle of claim 1 having a size of about 50 μm to about 5 mm.

16. 16. A composition for the localized delivery and sustained release of therapeutic cells and / or tissues and / or cell products at an implantation site, comprising a plurality of hydrogel microparticles according to any one of claims 1 to 15 dispersed or suspended in a biocompatible delivery vehicle.

17. 17. The composition of claim 16, further comprising a plurality of hydrogel microparticles comprising a three-dimensional matrix of a covalently crosslinked non-alginate based polymeric compound, the microparticles having a size greater than about 30 μm, and being essentially free of cells or tissue.

18. The composition of claim 16, wherein the plurality of hydrogel microparticles comprises a mixture of degradable and durable hydrogel microparticles dispersed or suspended in the biocompatible delivery vehicle.

19. 20. The composition of claim 18, wherein the durable hydrogel microparticles comprise insulin-producing cells and / or tissues and the degradable hydrogel microparticles comprise stem cells.

20. The composition of claim 16, wherein the plurality of hydrogel microparticles comprises a plurality of first hydrogel microparticles comprising a first cell type and / or tissue type and a plurality of second hydrogel microparticles comprising a second cell type and / or tissue type different from the first cell type and / or tissue type.

21. 21. The composition of claim 20, wherein the first cell and / or tissue type comprises insulin-producing cells and / or tissue and the second cell and / or tissue type comprises stem cells, or glucagon-producing cells and / or tissue.

22. The composition of claim 16, wherein the plurality of hydrogel microparticles comprises a plurality of first hydrogel microparticles having a first size and a plurality of second hydrogel microparticles having a second size different from the first size.

23. The composition of claim 16, wherein the vehicle is a biocompatible liquid suspension or gel.

24. 17. The composition of claim 16, wherein the medium is a cell culture solution or gel.

25. 17. The composition of claim 16, which is injectable or implantable via a catheter.

26. The composition of claim 16 comprising a therapeutically effective amount of the microparticles.

27. 17. The composition of claim 16 in the form of a putty, paste, or gel.

28. 17. The composition of claim 16 in the form of a liquid suspension.

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