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W2109655552 abstract "Stretched cavity-assisted molding of particles is a novel method that can produce monodisperse submicrometer hydrogel particles (see image) loaded with a model protein drug, fluorescein-isothiocyanate-conjugated bovine serum albumin. The method avoids agitation, solvents, and high-energy input by forming the particles inside preformed cavities. This method has the potential to be generalized to prepare multilayered or targeted particles containing fragile drugs for the pharmaceutical industry. Protein and DNA therapies are promising treatments that have been demonstrated in settings such as vaccination and insulin and hormone therapy, and will have increasing applications in disease treatment and cure.1–5 Drug-delivery systems (DDSs) are engineered to deliver these drugs better by taking into account in vivo conditions that might cause them to perform less effectively than demonstrated in vitro. The DDS serves to isolate fragile drugs from the body's internal conditions and immune functions, protect sensitive parts of the body from potent drugs, and enable the targeting and localization of drug effects at intended action sites in the body.6–10 One popular approach is to make micrometer- or submicrometer-scale particles loaded with the drug. 6–9,11 To make such particles with bio-macromolecular drugs, both top-down and bottom-up approaches developed for more stable chemical drugs have been investigated. Top-down approaches such as emulsification and aerosolization involve forming microscopic particles out of macroscopic quantities of bulk material.12–16 To reduce the particle size, high-energy mixing and organic solvent are often used, both of which have an undesirable denaturing effect on fragile biomacromolecular drugs. Bottom-up approaches,[17–21] such as block-copolymer self-assembly and liposome formation, can easily or spontaneously reach the submicrometer scale but are dependent on material properties to achieve self-assembly and good drug loading; potential drawbacks include possible cytotoxity of the carrier material or low drug loading. One recent top-down method using micromolding is particle replication in nonwetting templates (PRINT).22 PRINT reported monodisperse submicrometer particle sizes with flexibility in particle shape and composition, good biomacromolecular drug compatibility and efficient drug loading. Other unconventional techniques, such as continuous-flow lithography, have also been developed for high-throughput microparticle synthesis.23 Our novel method described here, stretched cavity-assisted molding of particles (SCAMP), is a novel top-down micromolding and nanomolding method designed to achieve monodisperse submicrometer size, minimal biomacromolecular drug destruction, high drug loading, and controllable particle shape. The SCAMP technique involves five steps (Figure 1): 1) An elastic polydimethylsiloxane (PDMS) mold with micrometer or submicrometer cavities is fabricated. 2) The mold is stretched and, while stretched, its cavities are filled with ultraviolet (UV)-curable precursor solution. The atmosphere around the mold is controlled by putting the mold inside an environment control chamber, purging the air, pressurizing with Ar gas to 9 bar (1 bar=100000 N m-2), and humidifying with wet tissue papers placed inside the chamber. 3) Excess precursor solution on the mold is removed by discontinuous dewetting, by tilting the mold24 so that the remaining liquid droplets inside the cavities are cured with 365-nm UV irradiation. 4) The cured droplets become solid particles and are partly ejected by releasing the tension on the mold outside the chamber. 5) The particles are forcibly removed from the mold by freeze-peeling, in which the particles are encased in ice and the elastic PDMS mold is peeled from them. Schematic presentation of SCAMP. a) PDMS rubber mold with microscale cavities, b) mold stretching, cavity filling (by vacuuming), and atmosphere conditioning inside chamber (O2 purging, 9 bar Ar, humidity provided by the wet tissue); c) discontinuous dewetting (by tilting of the chamber), and UV-curing to harden the precursor liquid droplets, d) partly ejected particles by releasing the stretch, e) complete detachment by freeze-peeling. We demonstrate SCAMP with three cavity dimensions: I) a 10-µm square cavity with 3.1-µm depth, II) a 2-µm circular cavity with 0.7 µm depth, and III) a 500-nm square cavity with 200-nm depth. The 10-μm particles produced from 10-μm cavities are considered too large for cell-targeted drug-delivery systems, but they were helpful for the eliminations of preliminary problems (e.g., to improve the efficiency of particle harvesting and water evaporation). Additionally, these larger particles are useful for understanding the particle ejection mechanism as they are clearly observable by optical microscopy. The 2-µm and 500-nm molds were used to make smaller particles that were relevant to DDSs and, therefore, demonstrate the applicability of SCAMP to drug delivery. To characterize the performance of SCAMP in terms of the incorporation of potential protein agents, fluorescent proteins, specifically green fluorescent protein (GFP) and fluorescein isothiocyanate-conjugated bovine serum albumin (FITC-BSA), were added to some particle precursor solutions to make fluorescent hydrogel formulations. This facilitates fluorescence microscopy determination of the distribution of the protein within the particles. Specifically, three particle precursor liquid formulations were used in the investigation. For preliminary investigations, formulation A, which consisting of 99/1 (w/w) polyethylene glycol diacrylate (PEGDA)/Irgacure 2959, was used. This formulation was also used as the resin component of the other two formulations. Drug loading was tested with a “B” formulation consisting of 70/30 (w/w) protein solution/formulation A. The protein solution is a GFP solution containing 8 mg GFP per mL deionized water (DI). BSA protein solution containing 5 wt% FITC-BSA in PBS (pH=7.4) was also used for the 500-nm particles. GFP does not dissolve in PEGDA alone and high water content in the formulation B improved the internal protein distribution of hydrogel B formed. FITC-BSA is more compatible with PEGDA and a higher loading can be used. Water evaporation from the mold cavities was studied with formulation C that consisted of 30/70 (w/w) DI water/A. Figure 2 shows representative scanning electron microscopy (SEM) images of the 2-μm and 500-nm particles after the ejection stage (step d in Figure 1). All three particle sizes were successfully ejected from the cavities in the PDMS upon release of the mold tension. The particles were elongated since the cavities were stretched during particle hardening. When refering to the particles, the unstretched cavity dimensions are used. Therefore, the 2-μm particles actually measure 3.5 μm×1.3 μm×0.6 μm (thick), the 500-nm particles measure 950 nm×380 nm×175 nm, and the 10-μm particles measure 21.3 μm×7.9 μm×2.1 μm. SCAMP B formulation (using GFP) particles after stretch release on a PDMS mold. a) 2-μm particles, b) 500-nm particles with I) low and II) high magnification. After removing the stretching force from the PDMS rubber, the particles were only partially ejected from the cavities because one end of the particles often remained stuck to the mold surface. To complete the detachment of the particles from the mold, freeze-peeling was employed (step e in Figure 1). Figure 3a shows that the rubber mold was stripped clean after the freeze-peel step; this is important for potential high-throughput applications, which would benefit from reusable molds. Both the 2-μm and 500-nm particles were completely transferred to other substrates when the ice that contained the trapped particles melted. The particles tend to aggregate (Figure 3b) during evaporation of the water from the freeze-peeling. Demonstration of freeze-peel particle harvesting of B formulation (using GFP) particles, a) PDMS mold covered with partly ejected 2-μm particles before freeze-peel (I) and stripped clean after freeze-peel (II), b) clustering of 2-μm particles on glass (I) and 500-nm particles on silicon substrate (II), after freeze-peeling and evaporation of the water peel medium. The water volume within each microcavity is so small that it evaporates almost instantly in standard atmosphere. In SCAMP, this would happen immediately after discontinuous dewetting. In spite of the water evaporation, the particles could still be cured and formed so that GFP was trapped inside the cured particles. However, the GFP inside these low water content particles was aggregated. This is visible under fluorescence microscopy as separated green dots inside the particles. Figure 4a shows a fluorescence microscopy image for 10-µm particles processed under standard atmospheric conditions. To reduce aggregation of protein and DNA therapeutic agents, we built a high-pressure, high-humidity environment chamber to reduce water evaporation. The discontinuous dewetting and curing process was performed in a high-pressure (9 bar) and humid atmosphere inside the environmental chamber. The 10-µm and 2-µm particles showed relatively smoother surface GFP distributions when the environment chamber was used (Figure 4b(I–II)). The surface layer is enriched with the GFP protein which seemed to prefer to migrate to the air–solution interface. This surface migration is more obvious with the 2-μm than the 10-μm particles due to the magnification used and larger surface area. With 500-nm cavities, the B particles did not appear fluorescent when GFP was used because of the relatively low incorporation of GFP and insufficient microscope magnification. GFP in the bulk protein/PEGDA solution aggregates into submicrometer-scale micelles, which retarded the filling of the submicrometer cavities by fluorescent GFP. When we used FITC-BSA as the fluorescent protein for the 500-nm B particles, fluorescence can be clearly observed for these particles (Figure 4c) because the FITC-BSA/PEGDA was a true solution and the FITC-BSA was present in higher concentration (5 wt%) than the GFP (around 0.8 wt%). Effect of environment conditioning on B particles examined with fluorescent microscopy a) without atmosphere conditioning, 10-µm particles using GFP; b) with high pressure and high humidity, showing relatively smoother GFP distribution of 10-μm particles and (I) 2-µm particles (II); c) with high pressure and humidity atmosphere showing smooth FITC-BSA distribution in particles ejected from the cavities but still adhered to the mold surface examined by fluorescence microscopy with a 20-µm scale bar (I), and particles harvested on the glass slide and examined by fluorescent confocal microscopy with 5-µm scale bar (II). Some proteins can diffuse out of the protein/PEGDA hydrogel when it is submerged in water, for example, during the freeze-peel step. Figure 5 shows the timed release of FITC-BSA from a B-hydrogel film measuring 1 cm×1 cm by 200 µm (thick). After the photopolymerization, the crosslinked B-hydrogel film was immersed in approximately 20 mL DI water. Figure 5a shows a representative luminescence emission spectrum of FITC-BSA aqueous solution. The relative fluorescence intensities of the immersion medium as a function of immersion time of the FITC-BSA/B-hydrogel film are recorded in Figure 5b–d. With prolonged immersion time, the fluorescence intensity increases, indicating more FITC-BSA leakage from the hydrogel. Fluorescence emission spectra (normalized, 488-nm excitation) of a) a typical aqueous FITC-BSA solution and the immersion medium of the B hydrogel film after immersion for b) 0 h, c) 2 h, d) 8 h, and e) 28 h. To confirm the preservation of biological activities of entrapped proteins after the SCAMP process, we perform biotin binding experiments with avidin-containing hydrogel particles. Briefly, a formulation consisting of CY3-labeled avidin, formulation A, and DI water in the ratio 1:40:40 (wt) was used for molding 10 µm hydrogel particles from a PDMS mold. The cured particles in the PDMS cavities were exposed to a fluorescein-labeled biotin solution (2 mg fluorescein biotin dissolved in 10 mL 50/50 mixture of isopropyl alcohol/DI water) for 3 h. Then, the hydrogel particles were washed thoroughly with DI water and observed using a fluorescence microscope. Figure 6a–c shows the images of particles using transmission (with rhodamine), and FITC modes respectively. (These three pictures were taken at the same location and the matching marker is shown.) Co-localization of the red and green fluorescence images indicates that the FITC-biotin prefers binding to the CY3-avidin particles. In a control experiment, particles were fabricated using pure PEGDA, exposed to fluorescein-labeled biotin for 3 h, and thoroughly washed with DI water. The fluorescence microscopy images show no binding of biotin to the CY3-avidin particles (data not shown). These results suggest that the biological avidin/biotin recognition is still preserved after our SCAMP process, confirming the preservation of the biological activity of the entrapped CY3-avidin protein. Fluorescence microscopy images to show the binding of FITC-biotin to CY3-avidin entrapped in 10-µm hydrogel particles in a) transmission mode, b) rhodamine mode, and c) FTIC mode. Even smaller size hydrogel/protein particles are achievable using smaller mold cavities, which may be produced by reported nanomachining or nanoreplication techniques. The SCAMP technique preserves the protein/DNA drug function by avoiding high-energy processing and organic solvents and is able to employ extremely aqueous precursor formulations. The technique is able to achieve monodisperse submicrometer particle sizes and uniform model protein drug loading. Drug encapsulation and timed controlled release are achieved. The PEGDA hydrogel has been reported to have a mesh size smaller than the dimensions of typical protein molecules.25 The diffusion-based release of the protein can be regulated by controlling crosslinking density or degradation of the polymer. The crosslinking material used here is PEGDA, which is biocompatible.14 The molecular weight of PEGDA can be varied from a few hundred to about 10000 Da to allow kidney clearance of the degraded fragments. SCAMP can also use other photocrosslinkable polymers as long as the resulting precursor formulation can discontinuously dewet the mold surface.24 A solution using high-molecular-weight dextran (20 wt%, MW = 450000 Da) in PBS cannot dewet the PDMS surface. The compatibility of the polymer and protein is also important. When polymalic acid was used instead of PEGDA for encapsulating the BSA, nonhomogeneity and phase separation were observed even while at 1000 rpm for 1 day. On the contrary, when a cationic chitosan derivative was used, gelation was observed quite soon. Stealth effect and specific targeting can be achieved by modification of the particle surfaces, presuming availability of residual acrylate end groups for crosslinking at the surface. Such modification could take place on the partially ejected particles before their removal from the mold, because most of their surfaces are exposed, or on the detached particles after the freeze-peel removal step. The ejection of the particles from the detensioned PDMS mold (prior to freeze-peel) is postulated to occur in the following sequence: 1) particle edges along the stretch direction break away from the cavity walls (the cavities widen during detensioning due to Poisson’s effect), leaving edges perpendicular to the stress direction still adhering to the cavities, 2) particles buckle as the cavities return to the unstretched state, and 3) one of the ends of the particles perpendicular to the strain direction breaks away from the cavities, allowing the particles to straighten half inside the cavities to give partly ejected particles. This is shown in Figure 7a(I). An SEM image of a particle still trapped in a cavity in the quite rare buckled configuration is shown in Figure 7a(II). A practical implication is that particle geometries with higher buckling tendency (slender, thin, and long particles) will be more easily ejected. Pressurized air spray (7 bar) and ultrasonication were unable to fully dislodge the partially ejected particles. Two-axis tensioning can completely detach the particles from the cavities (Figure 7b) but the stretching equipment would be significantly more difficult to build because a much larger rubber mold and larger forces are needed for the grippers. Therefore, harvesting by the relatively simple freeze-peel method was implemented (step e in Figure 1). SEM images of 10-μm PEGDA particles on a PDMS mold detached by a) pre-stretching; I) overview, II) a unique buckled particle; b) two-axis stretching performed by pre- and postcure stretching on orthogonal axes with a 1D tensioning device. (Dotted arrow, procure-stretch direction; solid arrow, postcure-stretch direction.) To evaluate the effectiveness of water-evaporation prevention in a conditioned atmosphere (high pressure and humidity), SEM images were made of 10-μm particles made with and without environmental conditioning. Representative SEM images of the cross-section of 10-µm C particles (Figure 8) show that particles cured in a conditioned atmosphere have a slightly bulging top surface (Figure 8a(I)), while unconditioned particles have a depressed top surface (Figure 8a(II)). Measurements of the particle volume in a sample of 30 particles cured in a conditioned atmosphere and 30 particles without conditioning show that the volume difference between the two types of particle is statistically significant and imply that the conditioned particles have 16.5% water by volume. The water content was originally 30% of the total weight, so it seems that the water-evaporation prevention is not 100% effective. An indication of the positive effect of conditioning is the relatively smooth GFP distribution on the surface of the 2-µm particles. If all water had evaporated prior to UV curing, even with high pressure and humidity conditions, the resulting particles would be expected to exhibit extensive GFP surface aggregation. Cross-section of 10-µm hydrogel particles (using 5-μm-deep cavities) a) with (I) and without (II) conditioned atmosphere and b) partial cavity filling due to water evaporation. One unexplored possibility with this method is the production of multilayered particles. Because the formation of the particles utilized cavities, layer-by-layer molding might be possible. One way to do this would be to use a polymer solution with high water content in a dry atmosphere for the dewetting step. Water evaporation prior to curing would result in partial filling of mold cavities with cured material (Figure 8b), possibly allowing other layers of material to be deposited in the cavities. One possible application is combination of materials/drugs that are incompatible with each other in the liquid form, and so cannot be mixed into a single solution. The PRINT technique lacks cavities and cannot form multilayer particles in this way. Both PRINT and SCAMP highlight the need for methods that can achieve a combination of various desirable factors and that are general enough to have the potential for further development, so that other requirements (e.g., stealth and targeting) can be addressed to achieve an effective DDS. We have demonstrated that SCAMP can make individual submicrometer-scale (950 nm×380 nm×175 nm) hydrogel particles encapsulating a biomacromolecular functional protein. This alternative method involves gentle processing conditions that will not destroy or denature biomacromolecular drugs. The premade cavities can have nanoscale dimensions and permit the formation of small particles without organic solvents or the high-energy agitation needed to finely divide agents with emulsification methods. The use of an aqueous solution for cavity filling permits high loading of water-soluble protein drugs. The protein-loaded particles are monodisperse, submicrometer-sized, and have high water content for high biomacromolecular drug loading. Other shapes and materials can also be used with this technique. Subsequent simple processing steps to coat the particles offer the potential to generalize this for the preparation of multilayered or targeted drug-delivery particles. Materials, chemicals, and molds: Poly(ethylene glycol diacrylate) (PEGDA) H2C=CHCO(OCH2CH2)nO2CCH=CH2 with average molecular weight (Mn) of 700 Da was purchased from Aldrich Chem. Co. Inc., USA. Sylgard 184 silicone rubber kit was purchased from Dow Corning Corp, USA. 4-(2-hydroxyethoxy phenyl-(2-propyl)ketone (Irgacure 2959), a photoinitiator, was purchased from Ciba Chemicals. FITC-BSA (fluorescein isothiocyanate-conjugated bovine serum albumin), CY3-avidin (ExtrAvidin-Cy3 conjugate) and FITC-biotin (biotin-4-fluorescein) were obtained from Sigma and used as received. Green fluorescent protein solution (2 mg mL-1) was synthesized in-house. All materials were used as received except GFP, which was concentrated to 8 mg mL-1. Three PDMS molds were used. The 10-μm PDMS mold was patterned with cavities 10-μm square and 3.1 µm deep, the 2-μm PDMS mold had round cavities of 2-µm diameter and 0.7-μm depth, and the 500-nm PDMS mold had cavities 500 nm square and 0.2-μm deep. These PDMS molds were replicated from polyurethane diacrylate second-generation molds that were themselves replicated from Si master molds patterned by deep reactive ion etching according to our previously reported procedures.26 Equipment: The following equipment was employed: I) an Axiovert 200M optical microscope with Axiovision 4.0 digital image capture software by Carl Zeiss Vision GmbH, II) a Zeiss LSM 510 meta confocal laser scanning microscope, III) a fluorescence spectroscope (iHR320 imaging spectrometer, Horiba Jobin YVON), IV) JEOL JSM-5600LV scanning electron microscope (SEM), and V) an Oriel flood UV-exposure source (model 97437) with Hg lamp (6 inch×6 inch beam size) and uniform intensity of 22 mW cm-2 in the 350–450-nm range (16 mW cm-2 measured at 365 nm). A uniaxis mold-stretching jig (hereafter called the stretcher) and environment control chamber with UV-window (poly(methyl methacrylate)) were designed and fabricated in-house; the stretcher enables stretching of the PDMS and the environmental control chamber enables evacuation, pressurization and gas type control in the environment. Procedure: To maintain the stretching of the mold throughout the cavity filling and UV-curing process, the stretcher was designed and fabricated. The stretcher grips the rubber mold tightly and uniformly applies a tensional load to stretch it in one dimension. Typically strains of nearly 100% were induced in the molds. After stretching, the molds were placed into the homemade environmental control chamber. The environment control chamber had a UV-transparent window and maintained vacuum and pressurized conditions. This chamber was used in step b of the process (Figure 1). For moldings under humidified conditions, wet tissue paper was placed inside the chamber before it was closed. The tissue provided high surface area for evaporation to humidify the atmosphere inside. To remove O2 and air, a cycle of evacuation, backfilling with Ar gas to 2 bar, and a pause period of 30 s was repeated 3 times. This also degassed the air bubbles that may have been trapped inside the cavities and precursor solution and ensured cavity filling. Afterwards, Ar pressure was increased to 9 bar; the high pressure retarded the water evaporation from the precursor solution. The chamber was then tilted to approximately 45° to allow the liquid to drain slowly. In step c (Figure 1), the formulation was UV-irradiated to harden it. The chamber was then depressurized and opened to remove the stretched mold. Detensioning the stretcher relaxed the mold and its cavities returned to their original shape. The resulting compression forces on the particles partially ejected them from their cavities (step d in Figure 1). In step e, freeze-peel was employed to completely detach the particles into a liquid medium such as water. The main idea of freeze-peeling is to trap the particles in another solid that allows the rubber mold to be peeled from them. The freeze-peel material must be easily switched between solid and liquid states so that subsequent collection of the particles is a simple matter of thawing (and evaporation of) the frozen material; water was therefore used. To successfully peel the particles, the ice must fully cover the top surface of the mold where the particles reside. Peeling of the rubber mold from the ice block should be done without shearing action, which would tend to dislodge the particles from the ice." @default.
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W2109655552 title "Stretched Cavity-Assisted Molding of Micrometer and Submicrometer Photopolymerized Hydrogel Particles" @default.
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