Sirolimus microspheres and method for producing sirolimus microspheres

Abstract

A method for producing monodisperse, homogeneous microspheres containing sirolimus is disclosed. The method may include producing sirolimus droplets using microfluidic step emulsification. The sirolimus droplets may include a drug loading wt% of less than about 50% or less than about 100%. The method may further include washing the droplets to remove poly(vinyl alcohol). The method may also include removing the solvent from the sirolimus droplets.

Description

[Technical field]

[0001] (CROSS REFERENCE TO RELATED APPLICATIONS) All applications for which a foreign or domestic priority claim is identified in an Application Data Sheet filed with this application are hereby incorporated by reference pursuant to 37 CFR 1.57.

[0002] The present disclosure relates to the field of microspheres produced by step microfluidic emulsification. [Background technology]

[0003] An implantable medical device is any device used for diagnostic and / or therapeutic purposes, such as stents, temporary or permanent indwelling catheters or balloons, that is introduced into the human body by clinical intervention and is intended to establish short-term contact with living tissue or to remain in place for a long period of time. Although these devices are subject to strict health and safety requirements, close and prolonged contact with the human body may increase the risk of adverse events and reactions, such as bacterial biofilm formation and restenosis. Blood vessels often suffer from recurrent restenosis after angioplasty. Restenosis is one of the significant risk factors limiting the long-term success of percutaneous coronary intervention.

[0004] Angioplasty balloons and coronary stents can be coated with anti-restenotic drugs, which are preferably released slowly after contact of the device with the arterial wall as part of angioplasty to inhibit neointimal hyperplasia, the primary cause of restenosis. Paclitaxel and sirolimus (rapamycin) and its derivatives are currently the two main anti-restenotic drugs used on or coated onto implantable drug-eluting medical devices to inhibit restenosis. Paclitaxel diffuses more readily through plaque and the arterial wall, but is less effective at inhibiting restenosis and has a narrower therapeutic window than sirolimus (SRL). As a result, sirolimus-eluting stents and sirolimus-coated angioplasty balloons are being developed.

[0005] Developments in drug-eluting technology include the adoption of drug carriers (excipients) that mix with the drug to form a coating and promote drug migration to the vessel wall. In the absence of excipients, the drug may form crystalline clumps on the device surface, which promotes washout and inhibits drug migration and absorption. This is especially true for drug-coated balloons, which have a short contact time between the inflated balloon and the vessel wall (30-120 seconds). Advantages of encapsulating the drug in carrier materials such as lipid excipients and / or biodegradable polymers include improved adhesion of the drug to the device surface, improved stability of the coating during handling, improved adhesion to the vessel wall during balloon expansion, and extended drug release to extend the therapeutic effect and minimize side effects such as dose-dependent toxicity. Coatings for drug-eluting medical devices often consist of an excipient layer that contains at least one therapeutic drug. Although initial drug concentrations are high with this approach, drug concentrations decline rapidly as the drug rapidly diffuses out of the surface region of the coating and the drug is rapidly depleted from the coating after reaching the vessel wall. One way to achieve a more uniform drug release over time is to prepare microparticles (sometimes spherical) consisting of a mixture of drug and carrier. These "microreservoirs" can be dispersed in a thin layer of carrier material to form a drug-releasing coating. After implantation, the drug molecules are slowly released from the microreservoirs deposited in the vessel wall by the expansion of the balloon, and each microreservoir acts as a miniaturized drug delivery system, releasing the drug into the secondary carrier layer. The drug is then diffused through the secondary carrier material to the living cells of the arterial wall or other tissue to be treated. The secondary carrier material should initially be one that is not easily washed off the balloon or catheter surface during advancement of the catheter to the treatment site, and then have the ability to be released and migrate to attach well to the endothelial lining of the artery, carrying the microreservoirs with it, thus providing a good geographical distribution of the microreservoirs on the treated tissue.

[0006] Polylactic-co-glycolic acid (PLGA) is the most widely used biomaterial for drug microencapsulation and is generally recognized as safe by the FDA and the European Medicines Agency (EMA). Currently, there are 19 FDA-approved PLGA-based drugs, which are mainly available commercially in the form of microspheres. Particle size is an important factor in their design, as it affects drug encapsulation efficiency, product injectability, syringe loading, biodistribution, and drug release rate. Conventional PLGA microsphere manufacturing methods, such as spray drying or emulsification using high-speed mixers or high-pressure homogenizers, followed by solvent evaporation, cannot precisely control the average particle size, and they show a wide particle size distribution with a coefficient of variation (CV) of about 30–50%. Particle size uniformity can be improved by expensive classification processes, but these processes result in high losses of drug and polymer. Although better control of particle size can be achieved using membrane emulsification, the CV is still relatively high, ranging from about 7% to about 20%.

[0007] Thus, there is a continuing need for manufacturing processes for drug-loaded particles with the key properties of 1-10 microns in diameter, uniform size and drug content, and inherent resistance to agglomeration for use in injectable and device coating applications. Agglomeration can be problematic because airborne particles have a tendency to adhere to each other to create larger, heavier aggregates.

[0008] Recent advances in microengineering and semiconductor technologies have enabled the fabrication of microfluidic chips to obtain monodisperse droplets with CVs below ∼3%. Droplet microfluidics offers many advantages over traditional emulsification methods, including unprecedented control over droplet size with no polymer / drug loss, high drug encapsulation efficiency as droplets are formed with negligible shear under continuous processing conditions, and the ability to operate in a closed environment enabling aseptic manufacturing to comply with CGMP regulations.

[0009] The main problem with conventional microfluidic devices, such as single T-junctions and ψ-junctions, is the low production rate, since droplets are formed one at a time. Moreover, the maximum throughput of droplets is limited by the transition from dripping to jetting, which must be avoided. Scaling up T-junctions and focusing nozzles is difficult because droplet formation in these geometries is controlled by the shear rate at the dispersed / continuous phase interface, which is sensitive to the fluid flow rate. As a result, small flow rate variations lead to large droplet size variations, making parallelization of these devices difficult. Step microfluidic emulsification is an alternative approach to generate uniform droplets in a low shear environment by exploiting the abrupt change in channel geometry from a shallow channel to a deep and wide reservoir. This mechanism was discovered in the mid-1990s but was largely ignored until recent years. The droplet size of a step emulsifier can be tuned solely by the internal channel geometry. The effect of fluid flow rate is negligible, making it easier to scale up individual microchannels. Summary of the Invention [Problem to be solved by the invention]

[0010] In the prior art methods, it has been difficult to formulate microspheres in the range of less than 20 microns in diameter with CVs in the range of less than about 3%. Disclosed herein is a method for making microreservoirs that are small in size (e.g., less than 20 microns) and uniform in composition. The amorphous composition of the small reservoirs as described herein is highly useful for mass production of drug delivery systems with precise pharmacokinetics that can be delivered and applied to micro-sized features, crevices, and tissue beds of the body using small, less traumatic device implants such as injectable slurries, stents, and catheters. The microreservoirs can optionally contain concentration-tunable polymeric components that can be selected to tailor the pharmacokinetics of the drug output of the microreservoir to treat various pathologies. The presently disclosed microreservoirs are useful for treating restenosis during or after angioplasty, injecting into the blood supply feeding tumors, injecting to reduce thrombosis in major arteries feeding major organs (e.g., brain, pancreas, liver), delivery in aerosols to treat conditions in the air-carrying structures of the lungs, delivery in ointments to treat skin conditions, delivery in injections and topical medications to treat degenerative eye conditions, delivery in oral medications, or delivery of a continuous flux of medication in the digestive tract, etc. In some embodiments, the disclosed microreservoirs can be used to deliver oral medications that include particles intended to reduce systemic inflammation throughout the body, which can lead to greater longevity. In some embodiments, the microreservoirs can be used for delivery of MRNA vaccines or DNA.

[0011] In some embodiments, disclosed is a method of producing aggregation-resistant monodisperse and homogeneous sirolimus microspheres. In some embodiments, the method includes using microfluidic step emulsification to produce droplets (sirolimus droplets) containing sirolimus, and the drug loading is about 100%. In some embodiments, the sirolimus microspheres further include poly(lactic-co-glycolic acid) (PLGA), and the drug loading is less than about 50 wt%. In some embodiments, the method includes washing the droplets to remove poly(vinyl alcohol). In some embodiments, the method includes removing the solvent from the sirolimus droplets to form particles of reduced size. For example, in some embodiments, the particles are reduced in size by the volume percentage of the solvent in the microsphere.

[0012] In some embodiments, a method of producing aggregation-resistant, monodisperse and homogenous sirolimus microspheres is disclosed. In some embodiments, the method includes producing sirolimus droplets using microfluidic step emulsification with a drug loading of less than about 50%. In some embodiments, the method includes washing the microspheres to remove poly(vinyl alcohol). In some embodiments, the method includes lyophilizing the sirolimus microspheres after solvent evaporation. In some embodiments, lyophilizing the sirolimus microspheres is configured to reduce particle size by about 10% to about 15%.

[0013] In some embodiments, the microfluidic chip used for microfluidic emulsification includes multiple junctions. In some examples, the multiple junctions of the microfluidic chip provide 3D flow focusing. In some embodiments, the microfluidic chip includes any number up to 10 junctions. In some embodiments, the microfluidic chip includes 7 junctions. In some embodiments, the microfluidic chip includes 10 parallel arrays of terraced microgrooves, the microgrooves having a depth between about 2 and about 5 μm. In some examples, the multiple parallel arrays of the microfluidic chip provide step emulsification.

[0014] In some embodiments, the surface of the microfluidic chip is washed with dichloromethane (DCM). In some embodiments, the microspheres are washed 6-7 times. In some embodiments, the microspheres are washed at least 8 times. In some embodiments, the droplets are formed using a drop method. In some instances, the drop method is used because it can generate highly uniform droplets. In some embodiments, the microspheres can also be formed using a squeezing region. In some embodiments, washing the microspheres further comprises centrifuging the suspension at 3500 rpm for about 10 minutes. In some embodiments, washing the microspheres further comprises adding an equal volume of about 0.05 wt% Tween 20 aqueous solution and vortexing for 5-10 seconds.

[0015] In some embodiments, the preparation of sirolimus droplets comprises using a continuous phase that does not contain isopropyl acetate (IPAc). In some examples, the microspheres can be loaded with a drug. Drug loading can be expressed as a weight percentage (wt%), where wt% is defined as the percentage of drug in the solid portion of the microsphere (i.e., the combination of drug and polymer). In some embodiments, the drug loading wt% of the sirolimus droplets is about 100%. In some embodiments, the sirolimus droplets may further comprise a polymer, and the drug loading may be about 30 wt% to about 100 wt%, about 30 wt% to about 80 wt%, or about 30 wt% to about 50 wt%. In some embodiments, the drug loading wt% of the sirolimus microspheres is about 33 wt%, about 34 wt%, about 35 wt%, about 36 wt%, about 37 wt%, about 38 wt%, about 39 wt%, about 40 wt%, about 41 wt%, about 42 wt%, about 43 wt%, about 44 wt%, between about 33 wt% and about 35 wt%, between about 35 wt% and about 37 wt%, between about 37 wt% and about 39 wt%, between about 39 wt% and about 41 wt%, between about 41 wt% and about 44 wt%, between about 33 wt% and about 42 wt%, between about 33 wt% and about 44 wt%, and any value between the recited ranges including the endpoints. In some instances, the drug loading of the sirolimus microspheres may be less than about 50 wt% or less than about 100 wt%. In some embodiments, removing residual solvent from the sirolimus droplets comprises lyophilization. In some instances, the majority of the solvent is removed by solvent evaporation, but the sirolimus droplets may also be lyophilized to remove any residual solvent. In some embodiments, the formed particles have a diameter greater than 8 μm. In some embodiments, the formed particles have a diameter between 1.8-8.4 μm. In some embodiments, the formed particles do not include Janus particles or phase separation. [Brief description of the drawings]

[0016] These and other features, aspects, and advantages are described below with reference to the drawings, which are for illustrative purposes and should not be construed as limiting the scope of the embodiments in any way. Moreover, various features of different disclosed embodiments can be combined to form additional embodiments that are part of this disclosure. In the drawings, like reference characters indicate corresponding features consistently throughout like embodiments. Each of the drawings will now be briefly described.

[0017] JPEG2025510567000001.jpg19117JPEG2025510567000002.jpg155117JPEG2025510567000003.jpg94117 DETAILED DESCRIPTION OF THE PREFERRED EMBODIMENTS

[0018] A system and method are disclosed for generating aggregation-resistant monodisperse microspheres containing sirolimus using step microfluidic emulsification and focused microfluidic conjugation. In some embodiments, the generated aggregation-resistant monodisperse sirolimus microspheres can then be used in drug-coated implantable devices and / or subcutaneous injections. In some embodiments, the microspheres may be substantially pure sirolimus. In some embodiments, the microspheres may further comprise a polymer. In some embodiments, the polymer may be polylactic-co-glycolic acid (PLGA) or polycaprolactone (PCL).

[0019] As described in more detail below, in some embodiments, the disclosed systems and methods generate aggregation-resistant monodisperse sirolimus microspheres with controlled size and internal morphology by microfluidic emulsification and subsequent solvent evaporation. In some embodiments, microfluidic emulsification can use step emulsification (FIGS. 1A-1E) and / or focusing conjugation (FIGS. 2 and 3A-3C). The systems and methods shown generate stable droplets in the drop-off regime that can be sustained for 48 hours, resulting in highly consistent size droplets created across all terraces of the microfluidic modules and chips used in step microfluidic emulsification. In some embodiments, the droplet size in the drop-off regime can be controlled by the smallest dimension of the droplet-forming microchannel (typically the microchannel depth). This confirms that droplet pinch-off can be driven by the capillary pressure gradient experienced by a deformed droplet squeezed in a shallow microchannel as it enters a deep microfluidic well.

[0020] In some examples, the size of the generated microspheres can be accurately predicted from the dispersed phase formulation and the size of the parent droplets. In some embodiments, aggregation-resistant monodisperse sirolimus (SRL) microspheres with diameters of 1.8-8.5 μm can be produced by microfluidic step emulsification and solvent evaporation using a single crystal silicon chip consisting of 10 parallel arrays of terraced microgrooves. In some examples, continuous generation of highly monodisperse (CV ≦ 3%, CV ≦ 4%, or CV ≦ 5%, etc.) droplets from microgrooves with depths of about 2-5 μm can be observed over 25 hours across all terraces. In some embodiments, the particle size after solvent evaporation can be reduced by about 10-15% during lyophilization without affecting particle size uniformity. In some examples, the microspheres can be small and highly monodisperse enough to be safely injected into the bloodstream using any size hypodermic needle and can be used as drug delivery coatings for implantable medical devices such as cardiac stents and balloon catheters.

[0021] Particle morphology may affect drug release rate, with less significant burst release expected from microspheres with more uniform internal structure. As currently disclosed, drug-containing microspheres may be substantially amorphous. In embodiments where the microspheres contain both drug and polymer, the degree of drug-polymer phase separation can be controlled by varying the drug content in the polymer matrix and the rate of solvent evaporation. In some embodiments, the polymer-drug phase separation becomes more pronounced as solvent evaporation is suppressed and drug loading is increased, resulting in heterogeneous patchy and patchy Janus particle morphology. As described in more detail below, accelerated drug release studies conducted at 37°C using a hydrotropic solution containing 5.8 M N,N-diethylnicotinamide revealed that an initial burst release released nearly one-quarter of the total drug.

[0022] As shown in the figures, the internal morphology of the drug-loaded microparticles was examined by FIB-SEM, FTIR, XRD, and fluorescence microscopy using Nile Red as a selective fluorescent stain with higher binding affinity for sirolimus compared to polyglycerol polyricinoleate (PGPR). In some embodiments, no new chemical bonds were identified in the FTIR spectra of drug-loaded microspheres, indicating the absence of covalent interactions between the drug and polymer during particle formation. The currently disclosed accelerated drug release test method showed that approximately 60% of the sirolimus was released within 24 hours, with the initial burst release consuming nearly one-quarter of the drug initially present in the microspheres. Furthermore, homogenous microspheres with complete mixing of drug and polymer can be produced when the drug loading is about 34% or less. In some embodiments, fast solvent evaporation / extraction can also lead to more homogenous particle production.

[0023] Further, a method for producing microspheres that contain only substantially amorphous drugs is disclosed.In some embodiments, the microspheres are substantially free of polymers.Such particles can be useful, for example, as a way to achieve faster dissolution of particles and provide higher maximum drug concentration when the particles are delivered to living organisms.In some instances, substantially polymer-free microspheres can be advantageous because they can minimize foreign body reaction to particles in tissues that are sensitive to polymeric materials and their decomposition by-products.

[0024] The currently disclosed method for producing drug-loaded monodisperse microparticles will enable the development of new, more effective implantable drug delivery devices and improved subcutaneous drug administration methods, leading to better therapeutics. In particular, the aggregation-resistant monodisperse sirolimus microspheres, with or without polymeric components, produced by microfluidic emulsification / solvent evaporation can open new routes for the fabrication of more effective implantable devices and the development of improved subcutaneous drug infusion methods for restenosis treatment. Similarly, the currently disclosed microfluidic approach for the reliable, robust, and scalable fabrication of highly monodisperse drug-loaded microparticles can be extended to other excipient / drug formulations, thereby improving the performance of many other subcutaneous drug therapies and implantable devices for controlled drug release.

[0025] (Microsphere Manufacturing System) The system including the microfluidic module and chip can be used to fabricate microspheres with monodisperse and controllable particle size. In some embodiments, stable droplet generation can be maintained for at least 24 hours, at least 36 hours, or at least 48 hours. Microfluidics can enable the manipulation of fluids at the micrometer scale. Advantages of microfluidics include the use of small amounts of reagents, miniaturization of devices, and low energy consumption. Microfluidics can also produce large quantities of droplets and / or particles that are uniform / monodisperse in size.

[0026] 1A-1E show one embodiment of a microfluidic module and chip 10. FIG. 1A shows a microfluidic rig 13a with two syringe pumps 11a and 11b, one for the continuous phase (11b) and one for the dispersed phase (11a). The pumps are fluidly connected to the module 13a such that the pump 11b for the continuous phase is in fluid communication with hole B (12b) and the pump 11a for the dispersed phase is in fluid communication with hole A (12a). In some embodiments, the microfluidic module may also include an emulsion collection syringe 11c. In some embodiments, the microfluidic module may also be equipped with a reflected light microscope 14 for observation of droplet generation. In some embodiments, the microfluidic module is made of stainless steel.

[0027] FIG. 1B illustrates an embodiment of a microfluidic chip 13b for use with the microfluidic module 13a. As illustrated, in some embodiments, the microfluidic chip 13b can include six dispersed phase channels 15a and ten parallel rows of terraced microchannels 15b labeled with terrace numbers 1-10. The chip of FIG. 1B can also include hole "A" which is an inlet hole for the dispersed phase 12a, hole "B" which provides an inlet hole for the continuous phase 12b, and hole "C" which provides an outlet hole for the emulsion 12c. In some embodiments, the dispersed phase channels 15a can be dead-end channels filled with the dispersed phase composition, and the continuous phase channels 16 are filled with the continuous phase composition. In some embodiments, the chip illustrated in FIG. 1B can have a width of about 25 mm. In some examples, the chip illustrated in FIG. 1B can have a height between about 25 mm and about 28 mm. In some examples, the chip can have a distance of about 3.5 mm between the centerlines of adjacent inlet holes for the continuous phase (i.e., hole "B"). In some embodiments, the chip may have a distance between the centerlines of the first and fifth inlet holes for the continuous phase (i.e., hole "B") of about 14.2 mm. In some examples, each of the cross-flow channels (i.e., the continuous phase channels) may have a length of about 11.1 mm. In some embodiments, the microfluidic chip may be made of single crystal silicon.

[0028] JPEG2025510567000004.jpg34117

[0029] FIG. 2 is a schematic diagram of another embodiment of a microfluidic module and chip 20. The microfluidic module can include a first syringe pump 21 with an aqueous phase (i.e., continuous phase) and a second pump 22 with an oil phase (i.e., dispersed phase). The first and second syringe pumps 21 and 22 deliver the continuous phase composition and the dispersed phase composition, respectively, to the microfluidic chip 23. A microfluidic junction 26 can be used on the microfluidic chip 23 to form an oil-in-water emulsion. The oil droplets formed in the aqueous phase can then be dried to form microparticles. The microfluidic module can also include a microscope 24 (optionally with a high-speed camera) for visualizing the formation of the microdroplets in real time. The microdroplets formed by the microfluidic module can then exit the microfluidic module where they can be collected 25.

[0030] In some embodiments, the microfluidic chip may be configured to include one or more droplet forming junctions 26. In some embodiments, the junction microfluidic chip includes a 3D flow focusing junction. As shown in FIG. 3A, the droplet forming junction 26 includes a dispersed phase channel 31, two continuous phase channels 32a and 32b, and an outlet channel 33 that come together to form the droplet forming junction. The dispersed phase channel 31 carries a dispersed phase (DP) solution toward the junction, and the continuous phase channels 32a and 32b each carry a continuous phase solution toward the junction. When these solutions meet at the junction, the dispersed phase is injected into the continuous phase flow, forming multiple droplets 34 surrounded by the continuous phase. FIGS. 3B and 3C show additional configurations of the droplet forming junction. In some embodiments, the microfluidic chip may comprise one single droplet forming junction. In some embodiments, the microfluidic chip may be a multi-junction chip. For example, the microfluidic chip may be comprised of 2, 3, 4, 5, 6, 7, 8, 9, or 10 junctions.

[0031] The system can provide for the production of drug-loaded microparticles. For example, the system can be used to form drug-loaded microspheres / microparticles by injecting a dispersed phase containing a drug and an organic solvent into a continuous phase containing deionized (DI) water and a surfactant. In some embodiments, the dispersed phase can further include a polymer or other carrier material.

[0032] In some embodiments, the design of the microfluidic chip can affect microparticle formation in addition to the emulsion formulation and solvent evaporation rate. For example, in the microfluidic chip shown in FIG. 1B, the flow rate ratio of the continuous phase to the dispersed phase can be greater than 1000. In some embodiments, the flow rate ratio of the continuous phase to the dispersed phase can be greater than 60, 100, 200, 300, 400, 500, 600, 700, 800, 900, or 1000. This can ensure that the droplets are generated in a large pool of continuous phase solution. For example, in an embodiment where dichloromethane (DCM) is used as the solvent for the dispersed phase, the solubility of DCM in water is about 1.60% at 20° C. When the flow rate ratio of the continuous phase to the dispersed phase is 62.5 or greater, the DCM is extracted into the surrounding aqueous phase without evaporation and is rapidly removed from the droplets in the chip. In some examples, the rapid removal of DCM from the droplets can result in a more homogenous structure of the drug-loaded particles than the slow removal of DCM by evaporation from particles stored in water. In other examples, such as the bonded microfluidic chip shown in Figures 3A-3C, the 3D geometry of the flow channels allows the dispersed phase jet to be completely surrounded by the continuous phase during droplet formation, which increases the contact area between DCM and water and therefore increases the removal rate of DCM.

[0033] (Fine particle manufacturing method) Drug-containing microspheres or microparticles can be produced using microfluidic step emulsification. A dispersed phase composition containing drug and solvent is injected into a continuous phase composition in a microfluidic chip to generate droplets containing the drug. The droplets are formed, suspended, and collected in an aqueous phase. In some embodiments, a collection syringe can be used to collect the droplets. In some embodiments, the solvent in the dispersed phase droplets is removed and extracted into the surrounding continuous phase (i.e., aqueous phase) solution. In some embodiments, the solvent further evaporates from the continuous phase solution into the air.

[0034] When the solvent in the droplets is removed, homogeneous drug-loaded particles are formed in the continuous phase solution. In some embodiments, the particles may be formed less than about 24 hours to about 48 hours after droplet generation. In some embodiments, the particles are further washed to remove the continuous phase solution before lyophilization. In some embodiments where polymers or excipients are present in the droplets / particles, phase separation may occur about 48 hours after droplet generation without a washing cycle, which leads to a lack of uniformity in the particles.

[0035] In some embodiments, the dispersed phase (i.e., oil phase) is composed of an organic phase solution. The organic phase solution can include a drug with an organic solvent. The drug can be, for example, sirolimus (SRL). In some embodiments, the organic solvent can be any water-immiscible solvent that can dissolve the drug. For example, the solvent for the SRL can include, but is not limited to, dichloromethane (DCM), acetone, chloroform, methanol, ethanol, ethyl acetate, acetonitrile, or isopropyl acetate. In some embodiments, the dispersed phase organic phase solution can also include a polymer, such as PLGA. In some examples, the continuous phase (i.e., aqueous phase) is composed of a cross-flowing aqueous solution. The aqueous solution can include, for example, deionized (DI) water and a surfactant. In some embodiments, the surfactant in the continuous phase can be PVA, poly(vinylpyrrolidone) (PVP), Tween 80, Triton X-100, sodium dodecyl sulfate (SDS), a Pluronic surfactant such as Pluronic F68 or Pluronic F127, or a monomethoxypolyethyleneoxide (MEPO)-b-PLA diblock copolymer. In some embodiments, the cross-flowing aqueous solution can include, for example, poly(vinyl alcohol) (PVA).

[0036] In some embodiments, the method of producing the microparticles described above can be used to encapsulate various drugs. In some embodiments, the drug can include any restenosis inhibitor. In some embodiments, the drug can include keoprofen (KET), ibuprofen (IBU), sirolimus, paclitaxel, or any of their derivatives. For example, sirolimus derivatives include, but are not limited to, everolimus, temsirolimus, pimecrolimus, ridaforolimus, umirolimus, biolimus A9, ABT578, zotarolimus, or any mTOR inhibitors called rapalogs (i.e., rapamycin and its analogs). The microparticles formed can be substantially pure drug-loaded microparticles, or PLGA and / or PCL microparticles.

[0037] In some examples, the microparticles can include various polymers, such as synthetic polyester-type polymers, such as poly(lactic acid) (PLA), poly(glycolic acid) (PGA), poly(lactic-co-glycolic acid) (PLGA) with different ratios of lactic acid and glycolic acid residues, polycaprolactone (PCL), poly(butylene succinate) (PBS), and the like.

[0038] In some embodiments, the microfluidic chips disclosed in Figures 1B and 3A-3C can be used to form microspheres of substantially pure sirolimus (i.e., substantially polymer-free). For example, an organic phase solution that is part of the dispersed phase composition is composed of sirolimus and a solvent and is injected through a microchannel into a deep well filled with a cross-flow aqueous solution of PVA as the continuous phase composition. In some embodiments, the solvent of the dispersed phase can be any solvent capable of dissolving sirolimus. For example, the solvent can be, but is not limited to, acetone, chloroform, methanol, ethanol, ethyl acetate, acetonitrile, or dichloromethane.

[0039] In some embodiments, microparticles can be formed using a junction microfluidic chip with a 3D flow-concentration junction (e.g., Figures 3A-3C). In some examples, the microfluidic chip can include a 3D designed flow-concentration orifice, which can result in faster solvent removal than a conventional flow-concentration chip with planar (2D) channels, since the continuous phase completely engulfs the dispersed phase jet during droplet generation.

[0040] In some embodiments, droplets can be generated in the bonded microfluidic chip under various regimes, as illustrated in Figures 4A-4D. Figure 4A illustrates droplet generation by squeezing 42, Figure 4B illustrates droplet generation by squeezing with a plug 43, Figure 4C illustrates droplet generation by dripping with a plug 43, and Figure 4D illustrates droplet generation by threading. In some embodiments, the plugs 43 are droplets with a diameter larger than the diameter of the outlet channel 41. In some embodiments, these droplets can have a non-spherical shape and occupy the entire cross section of the outlet channel 41. In some embodiments, the microfluidic chip can form a moving plug in the outlet channel 41.

[0041] In some embodiments, a cross-flow microfluidic chip with a microchannel array (as seen in Figures 1B-1D) can be used to scale up droplet / particle production. In some embodiments of the microfluidic chip, an organic phase solution (i.e., dispersed phase) can be composed of a drug (such as sirolimus) and an organic solvent (such as DCM), and optionally a polymer (such as PLGA), which is injected through a microchannel into a deep well (e.g., Figure 1D) filled with a cross-flowing aqueous PVA solution (i.e., continuous phase). In other embodiments, the continuous phase can be either an aqueous solution of poly(vinylpyrrolidone) (PVP), Tween 80, Tritox X-100, sodium dodecyl sulfate (SDS), Pluronic surfactants such as Pluronic F68 or Pluronic F127, or monomethoxypolyethyleneoxide (MPEO)-b-PLA diblock copolymer.

[0042] In some embodiments, the continuous phase may contain at least 1 wt% of a surfactant. In some embodiments, the continuous phase may contain from about 1 wt% to about 2 wt%, or from about 1 wt% to about 1.5 wt% of a surfactant. For example, the continuous phase may contain at least 1 wt% of PVA. In some embodiments, the continuous phase may contain from about 1 wt% to about 2 wt%, or from about 1 wt% to about 1.5 wt% of PVA.

[0043] As the droplets are generated in a large reservoir filled with the continuous phase solution, the solvent is rapidly extracted from the droplets into the surrounding aqueous phase, leaving behind homogeneous spherical microparticles, which leads to a more homogeneous drug-polymer particle structure.

[0044] Failure to wash the particles after droplet production can result in contamination of the final product with PVA. PVA is generally recognized as safe (GRAS) by the FDA, and the European Food Safety Authority (EFSA) has approved its use as a food additive, provided the total intake of PVA does not exceed 4.8 mg / kg bw / day. However, residual PVA may affect the bulk properties of the product. In some embodiments, the PVA concentration in the continuous phase can be minimized to reduce the initial level of PVA contamination and the number of washing cycles required. In some embodiments, the microparticles may be washed at least five times. In some embodiments, the microparticles may be washed six to ten times. In some embodiments, the microparticles may be further freeze-dried.

[0045] JPEG2025510567000005.jpg20117JPEG2025510567000006.jpg15117

[0046] JPEG2025510567000007.jpg23117

[0047] JPEG2025510567000008.jpg25117

[0048] JPEG2025510567000009.jpg10117

[0049] JPEG2025510567000010.jpg38117

[0050] In some embodiments, a clean hydrophilic surface of the silicon microfluidic chip can be advantageous after use (in some embodiments, the chip is reusable). In some embodiments, the clean silicon surface is hydrophilic due to the presence of polar silanol (-Si-OH) groups. In some examples, after emulsion production, the silicon walls can be covered with organic contaminants deposited during the production process. This can include, for example, PLGA, sirolimus, and PVA, which can result in a more hydrophobic surface that can impair droplet generation. In some embodiments, the organic contaminants can be partially removed by sonication of the chip with a household detergent solution. In some embodiments, chemical treatment of the chip can be performed after sonication, which can include oxygen or ozone plasma oxidation of the organic contaminants in a plasma reactor.

[0051] The hydrophilic chip surface can keep the contact angle between the organic droplets and the channel walls in the chip low. Because the step emulsification process is based on the Laplace pressure difference and the curvature imbalance along the droplet interface, uniform droplets can be generated in the chip continuously for at least 24 h by properly repairing the hydrophilic surface.

[0052] In some embodiments, the microfluidic chip can be cleaned and / or oven treated with DCM to remove residues before use or reuse. In some embodiments, about 90% of the residues can be removed after oven treatment. In some embodiments, the microfluidic chip can be cleaned by sonication of the microfluidic chip with a household detergent solution. In some embodiments, chemical treatment of the chip can be performed after sonication, which can include oxygen or ozone plasma oxidation of organic contaminants in a plasma reactor.

[0053] JPEG2025510567000011.jpg38117 Droplet size

[0054] In FIG. 4C, the length of the plug (L plug ) is the distance between the upstream and downstream ends of the plug. In some cases, the length of the jet (L jet ) can be the length of the thread connecting the newly formed droplets and the dispersed phase flow emerging from the orifice. In some embodiments, L plug and L jet can be controlled by the flow rate of the dispersed phase. In some embodiments, the higher the flow rate of the dispersed phase, the lower the L plug and L jet In some instances, the formation of plugs and jets is not recommended as this can lead to polydisperse droplets and particles.

[0055] JPEG2025510567000012.jpg48117

[0056] JPEG2025510567000013.jpg24117

[0057] JPEG2025510567000014.jpg36117

[0058] JPEG2025510567000015.jpg27117JPEG2025510567000016.jpg21117

[0059] In a cross-flow microfluidic chip with terraced microchannels, the microfluidic chip can be composed of two sets of cross-flow channels that supply the dispersed phase solution and the continuous phase solution. The two sets of cross-flow channels are connected by parallel stepped microchannels, where the dispersed phase solution is injected into the continuous phase solution to generate multiple droplets. The droplets are generated by a drop-by method in such a cross-flow microchip with terraced microchannels (MC).

[0060] JPEG2025510567000017.jpg66117

[0061] JPEG2025510567000018.jpg24117

[0062] (Drug Loading) In some embodiments, the SRL encapsulation efficiency in PLGA and PCL can be measured by dissolving the microparticles in acetonitrile (ACN) and measuring the dissolved particles using a UV / VIS spectrophotometer at wavelength (λ)=278 nm. In some embodiments, the encapsulation efficiency can be calculated as follows:

number

[0063] In some embodiments, the encapsulation efficiencies of PLGA and PCL were as follows:

number

[0064] (Washing of Microspheres) In some embodiments, after the microspheres are produced by step microfluidic emulsification, the solvent is removed from the microspheres into the surrounding aqueous phase. In some embodiments, once the solvent is removed, the microspheres are washed to form homogenous PLGA / SRL particles in an aqueous solution. In some embodiments, the microspheres can be washed between 24-48 hours after production. As described in more detail below, washing the microspheres can prevent the microspheres from undergoing phase separation. In some embodiments, the phase separation can be in the form of Janus PLGA / SRL particles in the aqueous phase. In some examples, the phase separation can be in the form of patchy Janus PLGA / SRL particles in the aqueous phase.

[0065] If the particles are not washed after droplet production, the formed microspheres may become contaminated with surfactants in the continuous phase, such as PVA. Although PVA is generally recognized as safe, residual PVA may affect the bulk properties of the microspheres. In some embodiments, the concentration of PVA in the continuous phase can be minimized to reduce the initial level of PVA contamination and the number of washing cycles required.

[0066] In some embodiments, washing the microspheres to remove PVA can affect particle interaction with cell membranes in the body by increasing the hydrophilicity of the surface. In some instances, the microspheres can be washed by centrifuging the emulsion. The supernatant of the centrifuged emulsion can then be removed and replaced with an equal volume of aqueous solution. In some embodiments, the emulsion can then be vortexed.

[0067] The amount of residual PVA adsorbed on the particle surface depends on the miscibility of the solvent blend with water. Particles prepared from solvent blends with higher amounts of isopropyl acetate (IPAc) are expected to have higher amounts of adsorbed PVA. In some instances, this may be due to the higher water solubility of IPAc compared to DCM. In some embodiments, when organic phase droplets can be formed in pure water, the formed PLGA particles can be hydrophobic due to the non-polar nature of PLGA. In some embodiments, when droplets are formed in a PVA solution, PVA can be adsorbed on the droplet surface such that the hydrophobic acetate tails of the PVA molecules can be immersed in the organic phase while the hydrophilic alcohol groups are exposed to the aqueous phase. In some embodiments, after solvent evaporation, a PVA coating remains on the particle surface, which can increase the hydrophilicity of the particles due to the exposed -OH groups on the surface. In some embodiments, the increased absorption of residual PVA increases the hydrophilicity of the particles.

[0068] (Solvent removal) In addition to washing the microspheres to prevent phase separation, the rate at which the solvent is removed can affect the amount of drug-polymer separation and whether there is uniform drug distribution within the polymer matrix. In some embodiments, the rate of solvent removal from the microspheres can be controlled by pre-saturating the continuous phase with IPAc.

[0069] The total specific volume of a solid polymer can be taken as the sum of the volume occupied by the polymer chains and the free volume of the material. When the organic solvent is rapidly removed from the droplets, the result is the formation of excess free volume trapped due to the rapid solvent removal, which can be reduced slowly by physical ripening or more rapidly by freeze-drying. In either case, the particle size after freeze-drying shrinks by about 10-15%, very close to the particle size predicted by the following equation:

number

[0070] JPEG2025510567000022.jpg31119JPEG2025510567000023.jpg15119

[0071] (Drug Release) In some embodiments, the duration of drug release in injectable, clinical long-acting PLGA-based depot formulations can range from one week to six months. Since this may be impractical in in vitro testing, the drug release rate can be enhanced by adding N,N-diethylnicotinamide (DENA). DENA has been shown to act as an effective hydrotrope (non-micellar solubilizer) for poorly water-soluble drugs. For example, the water solubility of paclitaxel from PLGA particles increases by several orders of magnitude in the presence of DENA. One explanation could be the result of faster degradation of PLGA in the presence of DENA due to hydrolysis of the ester bonds of the PLGA chains by base. Another explanation could be that this is caused by higher drug solubility due to non-stoichiometric accumulation of DENA around the drug. In some embodiments, long-acting PLGA-based depot formulations can exhibit an initial burst release that often releases a quarter of the total drug on the first day. As a result, drug concentrations on the first day of administration are often 100 times higher than steady-state drug concentrations in the blood. Similar behavior is observed where 242% of the encapsulated drug is released within the first 30 minutes. In some examples, the cause of the initial burst can be attributed to diffusion of drug molecules from the surface layer of the microparticles. The initial burst release can be followed by a steady-state release period of 30 minutes to 7 hours, during which the amount of SRL released increases to 24% to 29%. In some embodiments, after 24 hours, 59% of the drug still remains encapsulated in the particles and is released by quenching with acetonitrile (ACN). In some examples, during the steady-state release period, depletion of drug from the particles is offset by an increase in the permeability of the particle matrix due to DENA-induced PLGA degradation, resulting in a constant release rate of SRL. In some embodiments, the increase in the permeability of the polymer matrix is ​​a result of a decrease in the average molecular weight of the polymer and a decrease in the degree of entanglement of the polymer chains due to the hydrolysis of PLGA. According to the free volume theory of diffusion, the mobility of the PLGA chains can increase the permeability of the matrix to the SRL.

[0072] Working Example Example 1: Preparation of SRL-PLGA-microspheres The fabrication of aggregation-resistant monodisperse sirolimus-loaded PLGA microparticles by a step-microfluidic emulsification method is disclosed. The microparticles are fabricated with tunable size and internal morphology using a single crystal silicon chip with grooved microchannel arrays. The fabricated microparticles can be used for subcutaneous administration of sirolimus and as drug microreservoirs in sirolimus-coated balloons, providing new opportunities for the development of restenosis treatments and other pharmacological therapies based on subcutaneous injections and implantable drug delivery systems.

[0073] As discussed, microspheres can be generated using a microfluidic chip with terraced microchannels, where the microfluidic chip is composed of two sets of cross-flow channels that deliver the dispersed and continuous phase solutions. Three cross-flow silicon chips with terraced microchannels (MC) arranged in a 10 parallel array, as described in Table 1 below, can be used for droplet generation. Figure 1B shows an example of a 10 parallel array terraced microchannel microchip. Table 1 shows example dimensions of various microfluidic chips as shown in Figure 1E.

[0074] [Table 1]

[0075] Used chips can be cleaned in an ultrasonic bath set at 10% power using a mild detergent in a mixture of water and ethanol. The chips can then be rinsed with Milli-Q water, oven-dried, and transferred to a plasma cleaner to oxidize persistent organic contaminants that could not be removed by the detergent. In some instances, plasma oxidation is performed at 1×10 -5The cleaning process can be evaluated by measuring the contact angle between a water droplet and the dry chip surface. In some embodiments, the contact angle between a water droplet and a contaminated silicon surface is about 60°. After cleaning with dichloromethane (DCM), the contact angle was found to decrease to 32.4°, resulting from partial removal of organic contaminants. After plasma oxidation of the microfluidic chip, the contact angle further decreased to 16.9°.

[0076] Surface treatment using the aforementioned procedure was able to sustain droplet generation for at least 24 hours in all runs. The long-term stability of droplet generation on the CMS6-2 chip was compared in runs S2 and S5 in Table 2 provided below.

[0077] [Table 2]

[0078] In some embodiments, the average droplet size after 24 and 48 hours was the same (13.8 μm), but the CV increased slightly from about 2.5% after 24 hours to about 3% after 48 hours.

[0079] Figure 6 shows the temporal variation of droplet size generated from different terraces of the CMS6-1 chip described in Table 1. As shown in Figure 6, the variation in droplet size between different terraces is less than 1.5 μm, and the average droplet size of 23.5 μm is stable over time. Thus, about 89-95% of all channels can actively generate droplets at any time. Similarly, the droplet generation regime can transition from a dripping regime (i.e., small, uniform droplets) to a continuous flow regime (i.e., large, non-uniform droplets) at the critical dispersed phase velocity. In some embodiments, droplet generation may depend on the channel geometry, interfacial tension, and viscosity ratio between the two phases. In some examples, when triolein droplets were generated in about 1 wt% SDS aqueous solution, the critical velocity was 2.2 mm / s in a grooved channel of similar geometry. In the S1 run described in Table 2 below, the average dispersed phase velocity in the microchannel was 4.2 mm / s. In some embodiments, the critical velocity may be higher than 2.2 mm / sec because DCM is less viscous than triolein, and in some instances, the critical velocity may be higher for lower viscosities of the dispersed phase relative to the continuous phase.

[0080] JPEG2025510567000026.jpg57119

[0081] After producing microspheres by step microfluidic emulsification, the solvent is removed from the microspheres into the surrounding aqueous phase. As shown in Examples S5 and S6 of Table 2, in some examples, the PVA concentration in the aqueous phase can be reduced to about 1.5 wt% to about 1 wt% without a noticeable effect on the average droplet size and CV values. In some examples, reducing the PVA concentration below 1 wt% can have a detrimental effect on the droplet size uniformity.

[0082] Since PVA can affect the bulk properties of the product, the microspheres can be washed to remove the PVA. The emulsion was collected in an open syringe and the DVM was allowed to evaporate from the droplets to form particles. Residual PVA can then be removed from the suspension by multiple washing cycles. Each washing cycle consisted of centrifuging the suspension at 3500 rpm for 10 min, removing the supernatant with a Pasteur pipette, adding an equal volume of approximately 0.05 wt% Tween 20 in water, and vortexing for 5-10 s. After the final wash, the level of the supernatant in the vial was lowered to just below the level of the sediment, and the particles were dried using a benchtop freeze dryer. The contents of the vial were first frozen to approximately -20 °C and then allowed to dry by leaving it at a pressure of approximately 16 pa in the chamber and a condenser temperature of approximately -86.3 °C.

[0083] FIG. 9A shows an SEM image of SRL-loaded PLGA particles in an unwashed state. Unwashed particles may aggregate due to bonding by PVA films deposited between dried particles. Similarly, the formation of particle clusters by PVA bridges may change all particle size-dependent properties of the product, such as syringeability, dispersibility in secondary polymer matrices, and drug release behavior. FIGS. 9B-9C show SEM images of the same sample after eight washes. After eight washes, the particle surface is clean and smooth with no visible PVA film. In some embodiments, even after multiple washes, a small amount of PVA may remain adsorbed on the particle surface because PVA forms an interconnected network with PLGA. However, the remaining amount of molecularly bound PVA is too small to be quantified and does not affect the particle size distribution.

[0084] In addition to washing the microspheres to prevent phase separation, the rate at which the solvent is removed can affect the amount of drug-polymer separation and whether there is uniform drug distribution within the polymer matrix. The rate of solvent removal from the microspheres can be controlled by pre-saturating the continuous phase with IPAc. Figures 10A-10D and 11A-11D show the effect of organic solvent (IPAc) saturation of the continuous phase on the particle morphology of sample S1 (Table 2). Two different batches of sample S1 (S1A and S1B) were produced using PVA solutions with different solvent saturations. Figures 10A-10D show microspheres from sample S1A, and Figures 11A-11D show microspheres from sample S1B. In batch S1A, the continuous phase was a PVA solution at about 1.5 wt % pre-saturated with IPAc. In batch S1B, the continuous phase was a pure aqueous solution with the same PVA concentration, which allowed for an increased initial rate of organic solvent extraction from the droplets. Table 3 shown below shows the average particle size before and after lyophilization. The continuous phase was ∼1.5 wt % PVA saturated with IPAc (S1A) or a pure ∼1.5 wt % aqueous PVA solution (S1B).

[0085] [Table 3]

[0086] For both S1A and S1B, the particle size was approximately 7 μm or larger, but larger particles were formed in the S1B batch due to the faster solvent removal rate. Figures 10A-10B and 11A-11B show CLSM images of the particles before and after lyophilization. Figures 10C and 11C are schematic diagrams of the particle structure. Figures 10D and 11D are SEM images of particles cross-sectioned with a focused ion beam (FIB). The bright particle domains are enriched in sirolimus (SRL) and the dark particle domains are enriched in PLGA. Nile Red is preferentially absorbed by SRL over PLGA due to the high hydrophobicity of SRL with an octanol / water partition coefficient (log P) of 4.3. Nile Red is a hydrophobic compound and its fluorescence is strongly influenced by the polarity of the environment. As a result, the SRL-rich regions are less polar than the PLGA-rich regions.

[0087] As shown in Figures 10A-10D and 11A-11D, it is observed that the sirolimus is more uniformly dispersed in Figures 11A-11D (sample S1B) than in Figures 10A-10D (sample S1A) as a result of pre-saturating the continuous phase with IPAc. Figures 10B-10C show the formation of Janus particles with only one crescent-shaped SRL-rich region present on one side of the particle. The drug-polymer separation observed in Figures 10B-10C is consistent with the sirolimus loading of approximately 33% in sample S1, since the sirolimus region occupies a smaller particle volume than the PLGA-rich region (see column S1 in Table 2). Figure 10D shows that both regions contain both components due to incomplete phase separation. As shown in the FIB-SEM image in Figure 10D, the drug micropatches are embedded within the PLGA domains, and the SRL domains contain the PLGA patches. On the other hand, pre-saturating the continuous phase with IPAc would result in faster solvent removal from the droplets, suppressing drug-polymer separation and potentially resulting in incomplete separation of the SRL from the PLGA. Unlike Figures 10A-10D, no drug-polymer interface is observed in Figures 11A-11D. As shown in Figures 11A-11D, the faster the solvent removal rate, the more uniform the distribution of the drug in the solid matrix.

[0088] Example 2: Characterization of SRL-PLGA microspheres Table 2 above summarizes the emulsion formulations, fluid flow rates, droplet-particle sizes, and their coefficients of dispersion (CV) for the different samples.

[0089] As shown in Table 2 provided above, the long-term droplet generation stability in the CMS6-2 chip was compared in steps S2 and S5 of Table 2. As mentioned above, in some embodiments, the average droplet size after 24 and 48 hours was the same (13.8 μm), but the CV increased slightly from about 2.5% after 24 hours to about 3% after 48 hours.

[0090] In some embodiments, the average particle size produced from the systems disclosed herein can be between about 10 μm and about 1 μm for channel depths between about 6 μm and about 1 μm. In some examples, the average particle size is between about 1 μm and about 2.0 μm, between about 2 μm and about 3.0 μm, between about 3.0 μm and about 4.0 μm, between about 4.0 μm and about 5.0 μm, between about 5.0 μm and about 6.0 μm, between about 6.0 μm and about 7.0 μm, between about 7.0 μm and about 8.0 μm, between about 8.0 μm and about 9.0 μm, between about 9.0 μm and about 10.0 μm, and any value between these recited ranges, including the endpoints. In some embodiments, the average particle size can have a CV of particle size in the range of 1% to 5%. In some examples, the CV of particle size can range between about 1.0% and about 1.5%, between about 1.5% and about 2.0%, between about 2.0% and about 2.5%, between about 2.5% and about 3.0%, between about 3.0% and about 3.5%, between about 3.5% and about 4.0%, between about 4.0% and about 4.5%, between about 4.5% and about 5.0%, and any value between these ranges described inclusive of the endpoints. In some embodiments, the SRL-PLGA particles can be loaded with drug at a weight percent of about 30% to about 30%. In some embodiments, the SRL-PLGA can have a drug weight percent range between about 30% to about 35%, between about 35% to about 40%, between about 40% to about 45%, between about 45% to about 50%, and any value between these ranges described inclusive of the endpoints.

[0091] In the studies conducted and presented in Table 2, the average particle size of sample S1 was about 8.5 μm, with about 95% of the particles in the range of 7.8-9.2 μm. These particles can be injected through a 31-gauge needle (or smaller). In some embodiments, samples S2 and S3 may be suitable for intravenous delivery of sirolimus. As provided in Table 2, even the largest particles in the currently disclosed formulations are still several times smaller than the lumen of the smallest arteries (e.g., about 30 μm).

[0092] The average droplet size in the various chips was measured at regular time intervals over a period of 20-25 hours, and the results are shown in Figures 6-8. Figure 6 shows the average droplet diameter for the CMS6-1 chip at different terraces over a period of 25 hours. Figure 7 shows the average droplet diameter (bars) and coefficient of variation CV (solid line) for the CMS6-2 chip over a period of 24 hours. Figure 8 shows the average droplet diameter (bars) and CV (solid line) for the CMS6-3 chip over a period of 20 hours. As shown in Figure 6, the variation in droplet size between different terraces is below 1.5 μm, and the average droplet size of 23.5 μm is stable over time. Similar studies of the temporal stability of droplet size were also carried out in the CMS6-2 chip (shown in Figure 7) and CMS6-3 chip (shown in Figure 8). In some embodiments, the droplet size was lower than the CMS6-1 chip due to the smaller channel dimensions in the CMS6-2 and CMS6-3 chips. For the CMS6-2 chip, the 90% confidence interval for the mean droplet size was (13.8±0.6) μm with a CV=2.5%. In some embodiments, for the CMA6-3 chip (FIG. 8), the 90% confidence interval for the mean droplet size may be (5.2±0.3) μm with a CV=3.5%. In some instances, the CV values ​​slightly above 3% may be due to a larger error in estimating the size of the smaller droplets.

[0093] The average droplet diameter (μm) can be calculated for every junction on the MC chip (i.e., J1, J2, J3, J4, J5, J6, and J7) at 30 min, 120 min, 240 min, and 360 min. The dispersion coefficient can be calculated as follows:

number

[0094] The morphology of the particles can be visualized using confocal laser scanning microscopy (CLSM). A suspension of the microparticles or dry microparticles can be placed on a microscope slide and excited with an argon laser at 488 nm wavelength and a helium-neon laser at 543 nm wavelength. To enhance the observation of the polymer-drug distribution, the total emission can be split into two images and captured by two separate photomultiplier tubes (PMTs). PMT1 can capture the fluorescence at 515±30 nm (green region) and PMT2 can capture the fluorescence above 570 nm (yellow-red region).

[0095] Scanning electron microscopy (SEM) can be utilized to investigate the surface morphology of the particles and the efficiency of the developed cleaning procedure. In some embodiments, all micrographs can be taken with an aperture size of about 30 μm using a beam current of about 2.1 nA and an accelerating voltage of about 10 kV. If no crystals of PVA are found from the SEM images and there are no bridges of the PVA film between two particles, the cleaning can be considered successful. Particles with smooth and clean particle surfaces and no bridges between particles can be considered properly cleaned.

[0096] Imaging can be performed using a focused ion beam scanning electron microscope (FIB-SEM) combining ultra-high resolution SEM with precision FIB etching and deposition. The FIB acceleration voltage is about 30 kV and the ion beam current is about 20 nA. In some examples, the cross section can be cleaned at about 7 nA and a final clean can be performed at about 3 nA. In some embodiments, the current can be reduced to 30 pA to preserve the exposed cross section surface during imaging at 2 min / image.

[0097] X-ray diffraction (XRD) can be used to analyze the patterns of pure RL, blank PLGA particles, pure PLGA, and SRL-loaded PLGA particles (1:1 SRL:PLGA blend).

[0098] Attenuated total reflectance-Fourier transform infrared (ATR-FTIR) spectroscopy can be used to confirm the encapsulation of drugs into the produced microparticles. Approximately 2 to 3 mg of powder sample was placed on a universal diamond ATR top plate and observed at approximately 4000 to 400 cm -1 ATR-FTIR spectra can be recorded in the range of 100 s to 100 s. Depending on the sample, spectra can be acquired in approximately 32 seconds.

[0099] Example 3: Drug Loading FIG. 12 shows the effect of drug loading on particle morphology for particle sizes between 4.1 μm and 4.5 μm. As shown, the drug loading was (i) 33 wt% (S4), (ii) 42 wt% (S5), (iii) 44 wt% (S6), (iv) 45 wt% (S7), (v) 46 wt% (S8), (vi) 50 wt% (S9). The experimental conditions and emulsion formulations are shown in Table 2. The images in FIG. 12 were obtained using 20x (A) and 60x (B) magnification lenses. In some embodiments, both the drug and polymer exhibited fluorescence, but Nile Red exhibited higher adsorption when adsorbed to the drug molecules. As shown in FIG. 12, the drug-polymer distribution within the microparticles can be observed in the CLSM images, with the light areas corresponding to SRL and the dark areas corresponding to PLGA.

[0100] In some examples, drug segregation may depend on the drug loading. The extent of segregation ranges from homogeneous drug distribution at drug loadings of about 33 wt% and about 42 wt% (see (i)(A)-(i)(C) and (ii)(A)-(ii)(C)), to SRL patches embedded within a spherical PLGA matrix at drug loadings of about 44 wt% and about 45 wt% (see (iii)(A)-(iii)(C) and (iv)(A)-(iv)(C)), to a patchy Janus morphology at drug loadings of about 46 wt% and about 50 wt% (see (v)(A)-(v)(C) and (vi)(A)-(vi)(C)). In some embodiments, the patchy Janus morphology and the patchy Janus morphology are not stable during storage in aqueous phase, but gradually evolve into a fully segregated Janus morphology through Ostwald ripening. In some instances, the drug from the small patch dissolves into the surrounding aqueous phase and deposits into the larger patch. In some embodiments, this process may occur late in the solvent evaporation process due to inward diffusion of the SRL facilitated by residual solvent. In some embodiments, a similar patch coarsening process occurs in PCL-PLA composite particles, resulting in a variety of particle morphologies.

[0101] X-ray diffraction (XRD) analysis was performed on the freeze-dried S4 (see Table 2) microparticles to reveal the presence of impurities in the samples and to evaluate the crystallinity. The inclusion of SRL in PLGA can be confirmed by XRD analysis. The XRD pattern of the SRL-loaded PLGA microparticles does not show any sharp peaks, indicating that SRL is either molecularly dispersed in PLGA or present in the form of amorphous domains.

[0102] Fourier transform infrared spectroscopy (FTIR) analysis was performed on the sample S4 (see Table 2) microparticles. Figure 15 shows the FTIR spectra of pure PLGA, pure drug, and drug-loaded PLGA particles. The observed IR absorption peaks indicate the presence of characteristic bonds between PLGA and SRL. For example, the peak at approximately 1000 cm for SRL is -1 (1) and 1640 cm -1The peaks at (3) are due to the out-of-plane CH bending vibration and C=C stretching vibration of the -CH=CH- bond, respectively. These two bonds do not exist in PLGA, so the spectrum of PLGA did not show peaks at these frequencies. On the other hand, the spectrum of PLGA showed a peak at about 1100 cm -1 (2) and 1750cm -1 (4) Strong peaks were observed at approximately 3400 cm for SRL, which were due to the stretching vibrations of C-0-C and C=0 of the ester group, respectively. -1 The peak (5) is due to the 0-H stretching vibration. In PLGA polymers, the 0-H bond is present only in the end groups. No new peaks were found when ATR-FTIR was performed on the drug-loaded microparticles, since all peaks identified in the drug-loaded PLGA particles matched those of the individual components. This indicates that the drug did not react with the polymer during particle formation. Figure 15 also shows that peaks (1), (3), and (5) are negligible in the SRL-loaded PLGA particles, indicating that the SRL was not segregated to the particle surface but was homogeneously encapsulated within the particles. Figure 16 shows the FTIR patterns of SRL particles, PLGA particles, particles with a PLGA:SRL ratio of 4:1, and particles with a PLGA:SRL ratio of 1:4. As before, Figure 16 shows that no phase separation occurs in the homogeneous particles. Figure 16 shows that SRL was successfully encapsulated in PLGA and that particles with different SRL contents can be easily distinguished. FIG. 17 is another graph showing the FTIR patterns of SRL particles and particles with a PCL:SRL ratio of 4:1.

[0103] Example 4: Drug Release The release of SRL occurs much faster in dissolution media than in pure PBS. A burst release from PLGA particles occurs within the first hour, but then plateaus for a week. In contrast, 68–87% of the SRL was released from both PLGA and PCL particles after about 4 hours in dissolution media.

[0104] A 100 mL dissolution solution can be prepared in a volumetric flask by mixing 10 mL of 20 vol% polysorbate 20 (Tween 20), approximately 10 mL of 200 proof ethanol, approximately 25.9 mL of 5.8 M N,N-diethylnicotinamide (DENA) solution, approximately 44.1 mL of Milli-Q water, and approximately 10 mL of 10x phosphate buffered saline (PBS) for 1 hour at 37 °C. To measure the drug release profile of SRL, approximately 2 mg of lyophilized microspheres are placed in a microcentrifuge tube and 2 mL of dissolution solution is added. The tube is capped and alternately sonicated and vortexed for approximately 2 minutes to break up particle agglomerations and completely suspend individual particles in the solution. The tube is then placed in a shaker rotating at 250 rpm and incubated at 37 °C for approximately 25 minutes. The tube is then removed from the shaker and centrifuged at 13,000 rpm for approximately 5 minutes. The supernatant is extracted from the sample with a syringe and transferred to a labeled HPLC vial. The removed supernatant is replenished with fresh dissolution medium, and the same procedure can be repeated for samples taken after 1 hour, 2 hours, 3 hours, 4 hours, 5 hours, 7 hours, or 24 hours. After 24 hours, a quenched sample is prepared by adding about 2 mL of acetonitrile (ACN) to completely dissolve the drug-deprived particles and release the remaining drug. In some embodiments, all samples were stored at about -20°C before analysis. The concentration of SRL in the sample can be measured by high performance liquid chromatography (HPLC).

[0105] Example 5: Preparation of SRL microspheres Pure sirolimus microspheres were fabricated using a junction microfluidic chip containing a 3D flow focusing junction. The dispersed phase consisted of 96.5 wt% DCM with 3.5 wt% SIR dissolved in it. The continuous phase was 2 wt% PVA dissolved in deionized water. The solid particles were obtained from oil-in-water emulsion droplets collected after solvent evaporation. Briefly, the vial containing the collected droplets was left in a fume counter at room temperature to allow the DCM to evaporate. Indeed, DCM diffuses from the droplets into the surrounding aqueous phase, despite its low solubility in water (17.5 µg / L at 25 °C). Then, due to its high volatility (boiling point 39.6 °C), DCM evaporates from the air / liquid interface in the vial into the surrounding air. This gradual loss of solvent from the droplets increases the concentration of polymer and drug, eventually leading to the solidification of the droplets into particles. The particles were then washed with pure water to remove the PVA from the particle surface and frozen in a standard laboratory freezer at -20 °C. The majority of the volatile compounds (DCM and water) were then removed from the samples at a pressure of approximately 40 Pa and a condenser temperature of approximately -50 °C for 24 h. The samples were then reheated at 25 °C for 6 h to remove the remaining solvent.

[0106] As discussed for SRL-PLGA microspheres, successful droplet generation may depend on successful restoration of the clean hydrophilic surface of the silicon chip after use (in some embodiments, the chip is reusable). In some embodiments, organic contaminants may be partially removed by sonication of the chip with a household detergent solution. In some embodiments, sonication may be followed by chemical treatment of the chip, which may include oxygen or ozone plasma oxidation of organic contaminants in a plasma reactor. The hydrophilic chip surface may maintain a low contact angle between the organic droplets and the channel walls in the chip. Since the step emulsification process is based on Laplace pressure differences and curvature imbalances along the droplet interface, proper restoration of the hydrophilic surface may allow uniform droplet generation in the chip for at least 24 hours continuously.

[0107] In some embodiments, the continuous phase channel of the microfluidic chip can be cleaned by running soapy water and warm distilled water through the continuous phase channel. In some examples, warm distilled water can be run over the continuous phase channel at a high flow rate between 200 μL / min and 300 μL / min for 5 minutes. The dispersed phase channel of the microfluidic chip can be cleaned by running a pure solvent (e.g., DCM) through the dispersed phase channel. In some examples, the pure solvent can be run over the dispersed phase channel at a high flow rate between 200 μL / min and 300 μL / min for 5 minutes. In some embodiments, the microfluidic chip can also be deep cleaned with a 0.1 M to 0.5 M sodium hydroxide (NaOH) solution. This deep cleaning can be performed once a week, or at least once every two weeks, to ensure that the hydrophilic properties are maintained.

[0108] In some embodiments, used chips are cleaned in an ultrasonic bath set at 10% power using a mild detergent dissolved in a mixture of water and ethanol. The chips are then rinsed with Milli-Q water, oven dried, and transferred to a plasma cleaner to oxidize persistent organic contaminants that could not be removed by the detergent. In some examples, plasma oxidation is performed at 1×10 -5 The cleaning process can be evaluated by measuring the contact angle between a water droplet and the dry chip surface. In some embodiments, the contact angle between a water droplet and a contaminated silicon surface is about 60°. After cleaning with dichloromethane (DCM), the contact angle was found to decrease to 32.4°, resulting from partial removal of organic contaminants. After plasma oxidation of the microfluidic chip, the contact angle further decreased to 16.9°.

[0109] JPEG2025510567000029.jpg39119

[0110] In some instances, after the microspheres are produced by step microfluidic emulsification, the solvent is removed from the microspheres into the surrounding aqueous phase.

[0111] In some embodiments, washing the microspheres to remove PVA can affect particle interactions with cell membranes in the body by increasing the hydrophilicity of the surface. The amount of residual PVA adsorbed to the particle surface may depend on the miscibility of the solvent blend with water. Particles prepared from solvent blends with higher amounts of IPAc are expected to have higher amounts of adsorbed PVA. In some instances, this may be due to the higher water solubility of isopropyl acetate (IPAc) compared to DCM. In some embodiments, PVA can be adsorbed to the droplet surface such that when droplets are formed in the PVA solution, the hydrophobic acetate tails of the PVA molecules are immersed in the organic phase while the hydrophilic alcohol groups are exposed to the aqueous phase. In some embodiments, after solvent evaporation, the PVA coating remains on the particle surface, enhancing the hydrophilicity of the particles due to the exposed -OH groups on the surface. In some embodiments, the increased absorption of residual PVA can result in increased hydrophilicity of the particles.

[0112] Figure 13 shows the SEM images of microparticles with a PLGA:SRL ratio of 0:1. As shown, when the microparticles are 100% SRL, the microparticles formed are homogenous and have a regular spherical shape and are therefore completely amorphous. If the microparticles have a crystalline structure, they will have a non-spherical shape with sharp edges due to the regular arrangement of drug molecules within the crystal lattice.

[0113] In some embodiments, X-ray diffraction (XRD) analysis can be performed on the lyophilized microparticles to reveal the presence of impurities in the sample and to assess the degree of crystallinity. FIG. 14 is a graph comparing the XRD patterns of pure SRL, pure PLGA, 100% SRL particles, and 100% PLGA particles. As shown, pure SRL exhibited a crystallinity of about 50% to about 60%, pure PLGA exhibited a crystallinity of 0% to about 2%, 100% SRL particles exhibited a crystallinity of about 1% to about 2%, and 100% PLGA particles exhibited a crystallinity of 0% to 2%. The XRD spectrum of raw SRL powder shows sharp peaks at 2θ 6.8°, 9.8°, 12.2°, 14.5°, 15.9°, 19.9°, 20.2°, 21.5°, 22.5°, 23.3°, and 25°, indicating that the SIR is in a crystalline state. However, these sharp peaks were not observed in the XRD spectrum of pure SRL particles, suggesting an amorphous structure. These results suggest that the particle fabrication process caused a structural change of SRL from crystalline to amorphous. Fourier transform infrared spectroscopy (FTIR) analysis was performed on sample S4 (see Table 2) microparticles. Figure 16 is a graph showing the FTIR patterns of SRL particles, PLGA particles, particles with a PLGA:SRL ratio of 4:1, and particles with a PLGA:SRL ratio of 1:4. Figure 16 shows the successful encapsulation of SRL in PLGA and that particles with different SRL contents can be easily distinguished. Figure 17 shows the FTIR patterns of pure SRL particles and particles with a PCL:SRL ratio of 4:1.

Claims

1. 1. A method for producing aggregation-resistant, monodisperse, homogeneous sirolimus microspheres for coating implantable medical devices or for direct injection into biological tissue, the method comprising: producing microspheres comprising sirolimus using microfluidic step emulsification, said microfluidic step emulsification comprising injecting a dispersed phase composition comprising sirolimus and a solvent into a continuous phase composition; washing the microspheres to remove the continuous phase composition; and freeze-drying the microspheres.

2. The method of claim 1 , wherein the solvent consists of dichloromethane.

3. The method of claim 1 or 2, wherein the dispersed phase composition further comprises a polymer.

4. The method of claim 3 , wherein the polymer comprises PLGA.

5. The method of any one of claims 1 to 4, comprising freeze-drying the microspheres to reduce particle size by 10 to 15%.

6. The method according to any one of claims 1 to 6, wherein the microfluidic step emulsification uses a microfluidic chip comprising multiple 3D flow junctions.

7. The method of claim 6 , wherein the microfluidic chip comprises seven 3D flow junctions.

8. 7. The method of any of claims 1 to 6, wherein the microfluidic step emulsification uses a microfluidic chip comprising a cross-flow channel to deliver the dispersed phase composition and a parallel array of terraced microgrooves to deliver the continuous phase composition.

9. The method of claim 8, wherein the terraced micro-grooves have a depth between about 2 and about 5 μm.

10. The method of any of claims 8 to 9, wherein the flow ratio of the continuous phase composition to the dispersed phase composition is greater than 1000.

11. The method of any one of claims 8 to 10, wherein the microspheres are formed using a dropwise method.

12. The method of any of claims 1 to 11, wherein the formed microspheres have a diameter of less than about 8 μm.

13. The method of any of claims 1 to 12, wherein the formed microspheres have a diameter between about 1.8 and about 8.4 μm.

14. The method of any one of claims 1 to 13, wherein the microspheres are washed 6 to 7 times.

15. The method of any one of claims 1 to 14, wherein the microspheres are washed at least eight times.

16. The method of any one of claims 1 to 15, wherein washing the microspheres further comprises centrifuging a suspension containing the microspheres.

17. 17. The method of claim 16, wherein washing the microspheres further comprises adding an equal volume of about 0.05 wt% Tween 20 in water and vortexing for about 5 to about 10 seconds.

18. The method of any of claims 1 to 17, wherein the continuous phase composition does not include isopropyl acetate (IPAc).

19. 4. The method of claim 3, wherein the drug loading wt% of the dispersed phase composition is from about 33 wt% to about 44 wt%.

20. 20. The method of any of claims 1 to 19, wherein the solvent of the dispersed phase composition is selected from the group consisting of dichloromethane (DCM), acetone, chloroform, methanol, ethanol, ethyl acetate, acetonitrile, and isopropyl acetate.

21. The method of any of claims 1 to 20, wherein the continuous phase composition comprises deionized water and a surfactant.

22. 22. The method of any of claims 1 to 21, wherein the continuous phase composition is selected from the group consisting of polyvinyl alcohol (PVA), Tween 80, Triton X-100, sodium dodecyl sulfate (SDS), Pluronic surfactants such as Pluronic F68 or Pluronic F127, and monomethoxypolyethyleneoxide (MPEO)-b-PLA diblock copolymers.

23. The method of any one of claims 1 to 22, wherein the microspheres are washed with a 0.05% Tween 20 aqueous solution.

24. 1. A method for producing aggregation-resistant, monodisperse, homogeneous sirolimus microspheres, said method comprising: producing droplets comprising sirolimus using microfluidic step emulsification, the drug loading wt% being about 100%, the microfluidic step emulsification injecting a dispersed phase composition comprising the sirolimus and a solvent into a continuous phase composition; removing the solvent from the droplets to form the sirolimus microspheres; and washing the microspheres to remove the continuous phase composition.

25. 25. The method of claim 24, wherein the sirolimus microspheres are formed using a dropwise method.

26. The method according to any one of claims 24 to 25, wherein the microfluidic chip used for the microfluidic step emulsification comprises a plurality of junctions.

27. 27. The method of claim 26, wherein the microfluidic chip comprises seven junctions.

28. The method according to any of claims 26 to 27, wherein the microfluidic chip comprises 10 parallel arrays of terraced micro-grooves, the depth of the terraced micro-grooves being about 2 to about 5 μm.

29. The method of any of claims 24-28, wherein the formed sirolimus microspheres have a diameter of less than about 8 μm.

30. 30. The method of any of claims 24-29, wherein the formed sirolimus microspheres have a diameter between about 1.8 and about 8.4 μm.

31. The method of any of claims 24 to 30, wherein the sirolimus microspheres are washed 6 to 7 times.

32. The method of any of claims 24 to 31, wherein the sirolimus microspheres are washed at least eight times.

33. The method of any of claims 24 to 32, wherein washing the sirolimus microspheres further comprises centrifuging a suspension containing the sirolimus microspheres.

34. 34. The method of any of claims 32-33, wherein washing the sirolimus microspheres further comprises adding an equal volume of about 0.05 wt % Tween 20 in water and vortexing for about 5 to about 10 seconds.

35. The method of any of claims 24 to 34, wherein the continuous phase composition does not include isopropyl acetate (IPAc).

36. The method of any of claims 24 to 35, wherein removing the solvent from the sirolimus microspheres comprises lyophilization.

37. The method of any of claims 24 to 36, wherein the formed microspheres are substantially amorphous.

38. 38. The method of any of claims 24-37, wherein the dispersed phase composition solvent may comprise any one of dichloromethane (DCM), acetone, chloroform, methanol, ethanol, ethyl acetate, acetonitrile, and isopropyl acetate.

39. The method of any of claims 24 to 38, wherein the continuous phase composition comprises deionized water and a surfactant.

40. 40. The method of any of claims 24-39, wherein the continuous phase composition comprises polyvinyl alcohol (PVA), Tween 80, Triton X-100, sodium dodecyl sulfate (SDS), a Pluronic surfactant, such as Pluronic F68 or Pluronic F127, and monomethoxypolyethyleneoxide (MPEO)-b-PLA diblock copolymer.

41. The method of any of claims 24 to 40, wherein the sirolimus microspheres are washed with a 0.05% Tween 20 aqueous solution.

42. A method for producing highly uniform microspheres that contain a therapeutic agent in a substantially amorphous form.

43. 43. The method of claim 42, wherein the microspheres comprise a polymeric carrier material at a concentration of 0 to about 70%.

44. 44. The method of claim 43, wherein the polymer carrier is PLGA or polycaprolactone.

45. The method of any of claims 43-44, wherein the polymeric carrier and drug are molecularly dispersed and constitute a homogenous blend without distinct material domains.

Images