Langmuir 2008, 24, 9717-9726
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Evaluation of Continuous Flow Nanosphere Formation by Controlled Microfluidic Transport Bryan Laulicht,† Peter Cheifetz,† Edith Mathiowitz,†,‡ and Anubhav Tripathi*,‡ Department of Molecular Pharmacology, Physiology, and Biotechnology, and DiVision of Engineering and Medical Science, Brown UniVersity, ProVidence, Rhode Island 02912 ReceiVed March 25, 2008. ReVised Manuscript ReceiVed May 29, 2008 Improved size monodispersity of populations of polymer nanospheres is of enormous interest in the fields of nanotechnology and nanomedicine. As such, the knowledge of exact experimental conditions for precise production of nanospheres is needed for nonaqueous systems. This work presents the use of controlled microfluidic transport methods to study the experimental parameters for fabricating nanoparticles utilizing phase inversion. We report two microfluidic methods for forming polymer nanospheres in small batches to determine the formation conditions. These conditions were then implemented to perform higher throughput formation of polymer nanospheres of the desired size. The controlled microfluidic environment in the laminar flow regime produces improved size monodispersity, decreased average diameter, and affords a greater degree of control over the nanosphere size distribution without adding surfactants or additional solvents. Experiments show a nonlinear trend toward decreasing size with decreasing polymer concentration and a linear trend toward decreasing size with increasing flow rate indicating time-course-dependent nucleation and growth mechanism of formation for the resultant nanosphere population within the range of conditions tested.
Background and Motivation Nanospheres having diameters less than 1 µm offer significant advantages over more conventional microsphere formulations for oral drug delivery.1 Delivery of the entire drug-polymer complex, made possible by producing nanospheres on the order of the size of lipid rafts or clathrin coated pits has been shown to greatly improve cellular uptake over larger microspheres that are only uptaken by phagocytotic M-cells in the Peyer’s patches. Studies by the groups of Rejman2 and Florence3–5 indicate that polymer nanospheres differing by mere hundreds of nanometers in diameter experience widely different bioavailabilities and biodistributions within mammalian cells and among various tissues. Polymer nanospheres of approximately 250 nm in diameter remain in early stage endocytotic vesicles in cell culture; whereas larger particles of approximately 500 nm in diameter make their way to late-stage, enzyme containing vesicles. Jani et al.3–5 demonstrated a high degree of submicron diameter polymer sphere uptake in rats. In particular, spheres less than 1000 nm in diameter achieved gastrointestinal uptake of up to 34%.5 In the Jani et al. study, nanospheres with a mean diameter of 100 nm showed decreased uptake compared to 500 nm spheres indicating an active transcellular uptake mechanism for polymer nanospheres.5 Additionally, 100 nm diameter particles and smaller * To whom correspondence should be addressed. E-mail:
[email protected]. † Department of Molecular Pharmacology, Physiology, and Biotechnology. ‡ Division of Engineering and Medical Science. (1) Mathiowitz, E.; Jacob, J. S.; Jong, Y. S.; Carino, G. P.; Chickering, D. E.; Chaturvedi, P.; Santos, C. A.; Vijayaraghavan, K.; Montgomery, S.; Bassett, M.; Morrell, C. Biologically erodable microsphere as potential oral drug delivery system. Nature 1997, 386 (6623), 410-414. (2) Rejman, J.; Oberle, V.; Zuhorn, I. S.; Hoekstra, D. Size-dependent internalization of particles via the pathways of clathrin-and caveolae-mediated endocytosis. Biochem. J. 2004, 377, 159-169. (3) Florence, A. T.; Hillery, A. M.; Hussain, N.; Jani, P. U. Nanoparticles as Carriers for Oral Peptide Absorption - Studies on Particle Uptake and Fate. J. Controlled Release 1995, 36 (1-2), 39-46. (4) Jani, P.; Halbert, G. W.; Langridge, J.; Florence, A. T. The Uptake and Translocation of Latex Nanospheres and Microspheres after Oral-Administration to Rats. J. Pharm. Pharmacol. 1989, 41 (12), 809-&. (5) Jani, P.; Halbert, G. W.; Langridge, J.; Florence, A. T. Nanoparticle Uptake by the Rat Gastrointestinal Mucosa-Quantitation and Particle-Size Dependency. J. Pharm. Pharmacol. 1990, 42 (12), 821-826.
were found in the bone marrow, which indicates a level of tissue penetration that raises toxicity concerns for many drugs. Therefore the size range of greatest potential therapeutic benefit for polymer nanospheres in oral drug delivery is 200-1000 nm.3–5 The implications of these studies for nanomedical drug delivery technologies will be immense, necessitating the development of methods to produce monodisperse populations of nanospheres within the various size ranges. However, methods for entrapping sensitive therapeutics including proteins and biologics within polymer nanospheres are currently lacking in the literature and in commercial practice. Numerous microfluidic devices and techniques have been developed to produce polymer microspheres and microfibers6 that take advantage of emulsion formation,7,8 Rayleigh instability,7,9 photochemical cross-linking,10–12 and/or chemical synthesis.13 The size of the droplets and their corresponding microspheres produced by microfluidic emulsions utilize droplet breakup strategies to create monodisperse droplets on the micronscale. Monomers or prepolymers are introduced into catalysts and cross-linking agents in a controlled geometry yielding microspheres or microcapsules. In some setups reactions occur (6) Steinbacher, J. L.; McQuade, D. T. Polymer chemistry in flow: New polymers, beads, capsules, and fibers. J. Polym. Sci., Part A 2006, 44 (22), 6505-6533. (7) Ganan-Calvo, A. M. Polyphonic microfluidics. Nat. Phys. 2005, 1 (3), 139-140. (8) Whitesides, G. M. The origins and the future of microfluidics. Nature 2006, 442 (7101), 368-373. (9) Utada, A. S.; Lorenceau, E.; Link, D. R.; Kaplan, P. D.; Stone, H. A.; Weitz, D. A. Monodisperse double emulsions generated from a microcapillary device. Science 2005, 308 (5721), 537-541. (10) Dendukuri, D.; Pregibon, D. C.; Collins, J.; Hatton, T. A.; Doyle, P. S. Continuous-flow lithography for high-throughput microparticle synthesis. Nat. Mater. 2006, 5 (5), 365-369. (11) Dendukuri, D.; Tsoi, K.; Hatton, T. A.; Doyle, P. S. Controlled synthesis of nonspherical microparticles using microfluidics. Langmuir 2005, 21 (6), 2113-2116. (12) Nisisako, T.; Torii, T.; Takahashi, T.; Takizawa, Y. Synthesis of monodisperse bicolored janus particles with electrical anisotropy using a microfluidic co-flow system. AdV. Mater. 2006, 18 (9), 1152-+. (13) Sung, K. E.; Vanapalli, S. A.; Mukhija, D.; Mckay, H. A.; Millunchick, J. M.; Burns, M. A.; Solomon, M. J. Programmable fluidic production of microparticles with configurable anisotropy. J. Am. Chem. Soc. 2008, 130 (4), 1335-1340.
10.1021/la8009332 CCC: $40.75 2008 American Chemical Society Published on Web 08/06/2008
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at the interface between two flowing streams without the formation of an emulsion to produce microfibers by interfacial polymerization.6 Thermally or photoinitiated cross-linking of emulsified droplets can also be used to polymerize microspheres.10,11,14,15 Another photoinitiated cross-linking technique avoids the formation of droplets by using a photolithography-like setup for directly photopolymerizing a flowing polymer solution allowing for control over microparticle shape.12 Similarly, other aqueous microfluidic techniques8,9,16–21 have produced polymer microspheres; however, the production methods are limited to using an aqueous continuous phase. One promising method is the spontaneous phase inversion nanosphere formation (PIN) method22 in which nanosized particles of a chosen polymer can be prepared by pouring the polymeric organic solution (solvent) into another organic phase (nonsolvent) without any mechanical stirring. In contrast to previous polymer microsphere microfluidic studies, PIN does not involve the formation of droplets or cross-links. Instead, PIN utilizes the miscibility of the organic solvent and nonsolvent pair to enable production of thermoplastic polymer nanospheres on the nanoscale, at least an order of magnitude smaller than the microfluidic channels in which they are produced. As a result, PIN is a very promising method for producing polymer nanospheres from water-insoluble thermoplastic polymers, including those possessing desirable oral drug delivery properties including bioadhesive and bioerodible polymers. On the benchtop scale the PIN process produces nanosized particles of a chosen polymer prepared by pouring the polymeric organic solution (solvent) into another organic phase (nonsolvent) without any mechanical stirring or the creation of an oil/water interface in conventional glassware. For a microfluidic approach, due to the swelling of molded poly(dimethyl siloxane) (PDMS) and other polymer-involving systems the microfluidic platform had to be designed to withstand organic solvent usage. Materials including silicon, stainless steel, and glass were the most attractive options and glass was chosen due to the transparency, which allowed us to observe the flow pinching process, to confirm the lack of visible droplet formation, and to identify the nature of clogged channels when investigating the range of suitable polymer concentrations. Etched glass microchannels allowed for the introduction of chlorinated organic polymer solutions (e.g., PMMA in methylene chloride) into miscible organic polymer nonsolvents (e.g., (14) Dendukuri, D.; Gu, S. S.; Pregibon, D. C.; Hatton, T. A.; Doyle, P. S. Stop-flow lithography in a microfluidic device. Lab Chip 2007, 7 (7), 818828. (15) Dendukuri, D.; Hatton, T. A.; Doyle, P. S. Synthesis and self-assembly of amphiphilic polymeric microparticles. Langmuir 2007, 23 (8), 46694674. (16) Abraham, S.; Jeong, E. H.; Arakawa, T.; Shoji, S.; Kim, K. C.; Kim, I.; Go, J. S. Microfluidics assisted synthesis of well-defined spherical polymeric microcapsules and their utilization as potential encapsulants. Lab Chip 2006, 6 (6), 752-756. (17) Liu, K.; Ding, H. J.; Chen, Y.; Zhao, X. Z. Droplet-based synthetic method using microflow focusing and droplet fusion. Microfluidics Nanofluidics 2007, 3 (2), 239-243. (18) Liu, K.; Ding, H. J.; Liu, J.; Chen, Y.; Zhao, X. Z. Shape-controlled production of biodegradable calcium alginate gel microparticles using a novel microfluidic device. Langmuir 2006, 22 (22), 9453-9457. (19) Martin-Banderas, L.; Flores-Mosquera, M.; Riesco-Chueca, P.; RodriguezGil, A.; Cebolla, A.; Chavez, S.; Ganan-Calvo, A. M. Flow focusing: A versatile technology to produce size-controlled and specific-morphology microparticles. Small 2005, 1 (7), 688-692. (20) Okushima, S.; Nisisako, T.; Torii, T.; Higuchi, T. Controlled production of monodisperse double emulsions by two-step droplet breakup in microfluidic devices. Langmuir 2004, 20 (23), 9905-9908. (21) Seo, M.; Nie, Z. H.; Xu, S. Q.; Mok, M.; Lewis, P. C.; Graham, R.; Kumacheva, E. Continuous microfluidic reactors for polymer particles. Langmuir 2005, 21 (25), 11614-11622. (22) Carino, G. P.; Jacob, J. S.; Mathiowitz, E. Nanosphere based oral insulin delivery. J. Controlled Release 2000, 65 (1-2), 261-269.
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pentane) that induce phase separation of the polymer from the solvent-nonsolvent system to produce nanospheres. By choosing the appropriate solvent-nonsolvent pair that causes partitioning of the therapeutic agent with the polymer excipient, polymer encapsulated drug nanospheres are formed.1 On the macro-scale the formation of polymer nanoparticles by phase inversion leads to highly size polydisperse nanosphere populations when no excipients are added.1 Also, on the macro-scale, tuning of experimental parameters is very cost-intensive because it requires liters of organic solvents. Hence, the knowledge of the fundamental mechanism and exact experimental conditions for precise production of nanoparticles are still missing for organic solvent-based systems. In this paper, we describe a microfluidic phase inversion method for producing nanoparticles under highly controlled transport conditions using organic solvent-based system. The method requires only tens of microliters producing a tremendous cost savings. Although the microfluidic PIN procedure has a lower throughput than batch PIN, the methods presented provide increased control over production parameters such as flow rate, polymer concentration and dilution, greatly accelerating the pace of investigation into the mechanism of formation. Establishing the conditions leading to production of nanospheres in the desired size range for oral drug delivery on a microfluidic platform enables the rapid, low-cost investigation of processing parameters that lead to changes in the size distribution of nanosphere populations.
Experimental Methods Off-Chip Phase Inversion Nanosphere Formation (PIN). 100 µL of 0.0001 weight per volume percent 50 kDa PMMA (Mw/Mn ) 1.06) in methylene chloride (solvent phase) was ejected from a solvent-friendly pipet tip into 10 mL of pentane (nonsolvent). A 30 µL aliquot was then withdrawn from the bottom of the nonsolvent vessel by a fresh solvent-friendly micropipette tip and placed into an aluminum sample pan (Perkin-Elmer, Waltham MA). The liquid phase is allowed to evaporate and the resultant nanospheres were imaged by scanning electron microscopy (SEM). Experiments were repeated using starting concentrations of 0.001 and 0.01 weight per volume percent PMMA. Laboratory grade pentane and methylene chloride were supplied by Sigma-Aldrich and PMMA was supplied by Polymer Source (Montreal, Canada). Microfluidic Nanosphere Production with Flow Pinching. Microfluidic chips were fabricated in borosilicate glass substrate at the Brown University Microelectronics Facilities using a protocol based on standard microlithographic techniques. Briefly, the glass substrate was coated with chrome (∼800 Å thickness) and gold (∼400 Å thickness) using chemical vapor deposition after which a layer of Shipley 1818 photoresist was spin coated. After exposing the photoresist layer to UV through a negative mask, microchannels were etched in 49% hydrofluoric acid using calibrated etching versus time curves. A computer numeric control (CNC) lathe drilled holes in a second glass capping wafer, which later serve as reagent reservoirs. The etched glass wafer with the microchannel geometry is then bonded to the capping wafer using a controlled thermal bonding procedure.23–25 Figure 1 shows schematics and pictures of microfluidic chip used for nanosphere production. The chip has 4 reservoirs; each can hold up to 30 µL of solution, although experimentation was done with 20 µL of solution per reservoir. The channels are 75 µm wide and 12 µm deep. A Teflon caddy holds the glass chip against a metal edge backing plate, which applies uniform pressure to o-rings which (23) Kerby, M. B.; Lee, J.; Ziperstein, J.; Tripathi, A. Kinetic measurements of protein conformation in a microchip. Biotechnol. Prog. 2006, 22 (5), 1416-1425. (24) Kerby, M. B.; Legge, R. S.; Tripathi, A. Measurements of kinetic parameters in a microfluidic reactor. Anal. Chem. 2006, 78 (24), 8273-8280. (25) Lee, J.; Tripathi, A. Intrinsic viscosity of polymers and biopolymers measured by microchip. Anal. Chem. 2005, 77 (22), 7137-7147.
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Figure 1. (a) Full microchip assembly described from the top piece down: (i) pressure manifold with 8 threaded side ports for pressure lines. (A viton gasket to seal with the caddy is not shown.) (ii) Teflon caddy with through holes and O-ring seat. When assembled, the Teflon well has a capacity of 30 µL, while 6 µL is a functional minimum volume. Experiments were conducted with 20 µL volumes. (iii) Viton o-rings (iv) double layered, thermally bonded, glass microchip (wells and channels not shown). (v) threaded compression plate (vi) threaded microscope mounting plate. (b) Photograph of glass microfluidic cross chip containing a network of microchannels paired to microwells. Fluids move from reservoirs 1, 2, and 3 to reservoir 4, which serves as a collection reservoir for the resultant nanospheres. (c) Schematic of microfluidic flow pinching setup for nanosphere production. (d) A photograph of the microchannel flow showing 1:3 pinching. (e) phase contrast image of the branched polymer microstructure that forms upon phase inversion when the polymer concentration of the solvent is large (>0.01 wt %). (f) Schematic of the glass capillary tube setup for nanosphere production.
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seal the caddy and top glass plate. The pressure over the microfluidic chip reservoirs is controlled by a 4-port cartridge connected to corresponding pressure lines from the control system. Pressure control reduces the reagent requirement to a minimum of 6 µL/well and eliminates the holdup volume found with syringes and feed lines.23–25 The dilution and flow of the solvent and nonsolvent phases are regulated with a custom-designed two-component programmable control system. First, four independent pressure ports impose and measure the air pressure over any microchannel network containing fluids. Each pressure port has accuracy to within 0.01 psi of the assigned value over a (15 psi range and response time of 5 psi/s. Second, a custom LABVIEW software program and data acquisition boards (National Instruments Corporation) provide an automatic calibration and programming interface for the user. We have developed a simplified protocol for controlling the microfluidic chip dilutions by solving a system of momentum and continuity equations. In each channel i, the pressure drop can be related according to
∆Pi ) RiηQi
(1)
where η is the viscosity of liquid and Qi is the flow rate. The hydrodynamic resistance Ri of an isotropically etched microchannel is given by
Ri Ri ) RiRirect )
[
1-
∞
12 Li widi3
1 1 - exp(-nπwi ⁄ di) π wi n)1,3,5 n5 1 + exp(-nπwi ⁄ di)
192di 5
∑
]
(2)
where wi, di and Li are the width, depth and length of the microchannel i, respectively. The correction factor Ri, which is multiplied by the hydrodynamic resistance of rectangular channel, accounts for the isotropic shape of the channel. In the flow pinching experiment, reservoir 1 was filled with the polymer in solvent solution and the remaining three were filled with nonsolvent. The glass microchip is shown in schematic form and in a photograph in Figure 1a and b. A negative pressure of 1 psi was applied to reservoir 4 pulling the two nonsolvent and one solvent channels into the mixing channel inducing flow pinching. The solventnonsolvent interface has a sufficient difference in refractive index to allow viewing of the pinching flow using light microscopy shown diagrammatically in Figure 1c and in a light micrograph in Figure 1d. However, due to the miscibility of the solvent and nonsolvent, the interface is maintained only in the portion of the channel closest to the channel junction, beyond which mixing yields a single visually distinguishable liquid phase. Using the solvent trap setup, the viscosity of a 0.01% PMMA in methylene chloride solution was measured to be 0.4 cP, negligibly different from pure methylene chloride, on a TA Instruments AR2000 Rheometer. The viscosity of n-pentane used in calculations was measured to be 0.2 cP. The solvent phase flow rate was calculated using eqs 1 and 2 to be 0.093 nL/s. Flow pinching conditions produce a 30:70 dilution ratio of the solvent to nonsolvent phases at the junction. After ten minutes of run time, the contents of well 4 were collected for size analysis. Microfluidic PIN Nanosphere Production without Flow Pinching. Next we tested the second solvent/nonsolvent configuration. In this configuration, reservoirs 1, 2, and 3 (Figure 1a) were filled with the dilute polymer solutions and reservoir 4 was filled with the nonsolvent pentane. Negative pressure of 1 psi was applied to the nonsolvent reservoir causing flow of the dilute polymer solution into reservoir 4 via the cross chip microchannels avoiding the flowpinching phenomenon. In this configuration the solvent phase flow rate is calculated to be 0.31 nL/s. Capillary Tube PIN Nanosphere Production. In an effort to increase the scale of nanosphere production, glass capillary tubes (Labcraft 100 µL disposable glass micropipette tubes) were press-fit into male luer to 1/16 in. tube fittings (McMaster Carr) heated to
Figure 2. (a) Exemplary scanning electron micrograph (SEM) of PIN nanospheres produced (b) NIH ImageJ generated ellipsoid outlines generated during nanosphere population size measurements.
120 °C. The glass capillary tubes were interfaced with solventfriendly syringes that were filled with dilute 0.001 wt % PMMA in methylene chloride (Figure 1f). Each solvent-friendly syringe containing dilute polymer in organic solvent solutions was placed into a Harvard Apparatus Pump 11 Pico Plus syringe pump (Hamden, CT). The syringe pump was set to flow organic solvent through the glass capillary tube (inner diameter ) 1 mm) at rates ranging from 1-100 nL/s into a 10 mL Pyrex beaker containing 10 mL of nonsolvent, pentane. After formation, 50 µL of each sample are collected for size analysis. Nanosphere Sizing. For the microfluidic nanospheres production methods, at the end of each 10 min run, the entire contents of the collection reservoir were transferred by micropipette into an aluminum sample pan (Perkin-Elmer, Waltham, MA). Aluminum sample pans were used because of their geometry and conductivity, providing an ideal vessel for evaporating volatile organics leaving behind polymer nanospheres. Sample pans were placed on SEM stubs with double-sided carbon tape and sputter-coated with 50-100 Å of gold-palladium (Emitech K550, Kent, UK). The stub was inserted into the Hitachi S-2700 scanning electron microscope (Tokyo, Japan) with an accelerating voltage of 8 kV. The microscope was aligned and then digital pictures were obtained via the Quartz PCI digital imaging system and software (Quartz Imaging Corporation, Vancouver, BC). Resultant images were analyzed for Ferret’s mean diameter of fitted ellipsoids using NIH ImageJ (Bethesda,
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MD) as depicted in Figure 2 to determine Ferret’s mean diameter of the nanospheres. For the off chip and capillary tube nanospheres production methods 30 µL of the resultant nonsolvent bath collected from the bottoms of the beakers are transferred by micropipette into aluminum sample pans and the above sizing procedure is the same. Statistical Analysis. Statistical analysis was performed using SPSS software (Chicago, IL). In populations of nanospheres produced that had inhomogeneous variances, the Welch and Brown-Forsythe robust tests of equality of means were run followed by a Dunnett T3 posthoc test. The Student’s t test was used to compare nanosphere populations produced by 0.0001% PMMA solutions on both microfluidic methods since the variances were homogeneous. From the SPSS calculations we calculated p-values. The p-value measures consistency between nanosphere populations by calculating the probability of observing the same results between experimental data sets.
Results and Discussions We seek to understand the formation of PMMA (poly(methyl methacrylate)) chains into spheres. PMMA chains of molecular weight Mw ) 50 kDa (Mw/Mn ) 1.06) consisting of N statistically independent-segments each of length b (the “Kuhn length”). The number of segments and the “Kuhn length” in the equivalent freely jointed Kuhn chain are computed as N ) 3Mw sin2 θ/MoC∞ ) 130, and b ) C∞l/sin θ ) 1.72 nm, where C∞ ()9.1) for polyethylene oxide chains) is the characteristic ratio, l ()0.154 nm) is the carbon-carbon bond length, Mo ()100 g/mol) is the molecular mass of the repeat unit and θ ()54.5°) is the half angle between carbon-carbon bonds in a polymer chain. The rootmean-square end-to-end distance of the equivalent Kuhn length is
〈R2〉1⁄2 ) √Nb2 ⁄ 3 ) 11.33nm A solution in MeCl2 of such polymer chains of narrow molecular weight distribution with a concentration c per unit volume is injected. The number density of the chains can be computed as n ) cNA/Mw, where NA is Avogadro’s number. The average distances between chains for 0.01%, 0.001% and 0.0001% solutions are approximately 58 nm, 125 and 271 nm, respectively. Hence, the distance between the nucleation sites grow nonlinearly with concentration. Since the distances between chains are greater than the radius of gyration, the polymer solutions are in the perfectly dilute regime therefore polymer chains must diffuse, agglomerate, and collapse to result in nanoparticles of sizes observed in the experimental results. Results of four phase inversion nanosphere (PIN) production methods are reported. In the first method the PIN are formed in conventional way. Here, the polymer solution is put into a beaker full of nonsolvent. In the second method, PIN spheres are formed by a microfluidic flow pinching (Figure 1a). Here, the polymer molecules flow in contact with nonsolvent molecules and diffusion occurs across the microchannel laminar flow. In the third method, the solvent phase flows directly into the stagnant nonsolvent well without first pinching the flow. Here, the polymer molecules flow from microchannel into the pool of non solvent molecules. The diffusion mixing is similar to a radial “source” flow mixing. In the final method, the polymer molecules are injected into the pool of nonsolvent molecules using a capillary flow. The diffusional mixing is similar to a “fluid jet” mixing. In all cases the nanospheres generated were imaged and sizes were analyzed on NIH ImageJ software, an example of which is shown in Figure 2. Off Chip PIN Experiments. Our experimental runs using 0.0001 and 0.001 weight per volume percent PMMA solutions
Figure 3. (a) Scanning electron micrograph (SEM) of nanospheres produced off chip. 0.01 wt % PMMA was used. (b) Size histogram of nanospheres.
showed no visible formation of particles in the bulk phase. It appears that particles were either not formed or smaller than lowest detectable diameter by SEM (∼50 nm). At these very low concentrations, the polymers collapse as isolated coils that are very far apart. Experiments were then repeated using starting concentration of 0.01 weight per volume percent PMMA and synthesized spheres that were observed under the SEM shown in Figure 3. Here, polymer chains are in close enough proximity to attract each other while phase inverting to form the cores of polymer nanospheres, but polymer chains at the surface of the nanospheres contact pure solvent as it is driven from the core of the polymer due to the phase inversion coupled with diffusion into the nonsolvent. Owing to high cost of their surface energy, nanospheres would like to stick together, forming larger clusters with lower surface energy per molecule due to reduced surface area of contact with the nonsolvent. This tendency results in formation of bigger nanoparticles and agglomerates. The figure clearly shows signs of agglomerated nanoparticles of spherical and nonspherical shapes of different sizes. A size histogram is shown in Figure 3b. The data shows 660 ( 48 nm effective diameter nanoparticles including agglomerates. The data shows huge scatter in size and 53% of the nanosphere population is larger than 500 nm and 12% is larger than 1 µm. Moreover, the population of nanospheres produced by PIN not only is limited
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Figure 4. SEM of a population of nanospheres produced by glass chip microfluidics.
to the highest polymer concentration used on the glass microfluidic chips, but also has one of the largest coefficients of variance measured in testing, 52%, in which coefficient of variance is defined as the percent that the standard deviation is of the mean diameter. Hence, in the previous PIN investigations,1,22 as with the off-chip production, the uncontrolled introduction of the solvent into the nonsolvent phase perhaps lead to nonhomogenous nucleation sites resulting in increased polydispersivity and coefficient of variance. It should be noted that the off-chip experiments require large amounts of solvent and polymer solutions and hence it would require large number of experiments to obtain a desired relationship between PMMA concentration and average particle size. Microfluidic Nanosphere Production with and without Flow Pinching. We first investigated the effect of polymer concentration on particle production. Dilute polymer (PMMA) in methylene chloride (MeCl2) solutions were run in the above chip conditions at concentrations ranging from 0.0001 to 1 wt % in orders of magnitude. Above 0.01 wt % polymer formed network structures in the microchannels (75 µm wide and 12 µm deep) along the solvent/nonsolvent interface indicating that the polymer chains precipitated in close enough proximity to join into a bulk microstructure rather than discrete nanospheres as shown in Figure 1e. This experimental result suggests that the rapid collapse of polymer chains across the entire cross-section of the channel. The diffusion time for a 50 kDa polymer in a good solvent to travel across 75 µm wide channel is td ∼ w2/2D ) 752/2 · 67 ) 42 s. Here, D ) 67 µm2/s is the molecular diffusivity of PMMA.26 Since the diffusion of solvent and polymer molecules across the width of the channel was rapid, the time scale of this particle growth was almost instantaneous. The residence time of the polymer molecules while traversing that microchannel is tr ∼ la/Q ) 400 s in the flow-pinching and tr ∼ lA/Q ) 120 s in the nonflow pinching configurations. Therefore the residence time in the channel far exceeds the diffusion time. Finally, it is noted that the overlap concentration (c*) for PMMA in the solvent is c* ≈ 2.5/[η] ) 0.012 g/mL ≡ 1.2% Here, [η] is the measured intrinsic viscosity of 50 kDa PMMA in a MeCl2 solution. Microfluidic nanosphere populations (Figure 4) produced in glass chip microfluidics are significantly different from the offchip (Figure 1), uncontrolled nanosphere formation (p < 0.01). (26) Larson, R. G. The Structure and Rheology of Complex Fluids. Oxford UniVersity Press: New York 1999, xxxx, xxxx.
Figure 5. Size histograms of nanosphere populations produced by the flow pinching glass chip microfluidics methods.
In the microfluidic flow pinching configuration, 0.01 wt % 50 kDa PMMA (Mw/Mn ) 1.06) solution formed 731 ( 32.7 nm diameter nanospheres on average (Figure 5). At 0.001 and 0.00014 weight percent 533 ( 18.5 nm and 517.4 ( 19.6 nm diameter spheres respectively were produced (Figure 5). In the nonflow pinching conditions on the same microfluidic chips, similar results were achieved: 846 ( 19.3 nm at 0.01, 529 ( 23.7 nm at 0.001, and 525 ( 17.4 nm at 0.0001 PMMA weight percent (Figure 6). On the whole, in both cases mean nanosphere size decreased
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Figure 7. Mean nanosphere diameter plotted as a function of polymer concentration for both microfluidic nanosphere production methods. ** ) Statistical difference between methods, p < 0.01; † ) statistical difference between concentrations, p < 0.001.
Figure 6. Size histograms of nanosphere populations produced by glass chip microfluidics method without flow pinching.
significantly as polymer concentration decreased (Figure 7). However, in the flow pinching case the mean sphere volume decreased by a factor of 2.6 and without flow pinching the mean sphere volume decreased by a factor of 4.1. Moreover, the 10fold differences in initial polymer concentration do not linearly correlate with the mean nanospheres volume for either microfluidic technique possibly indicating a difference in degree of polymer chain collapse and/or that solvent is trapped within the
nanospheres. Note that the nanosphere sizes are much smaller than microchannel width (75 µm) or depth (12 µm). The size depends only on the local concentration and diffusion time of polymer chains in the microchannel. It should further be noted that the synthesis techniques reported in literature only demonstrate particle formation of sizes comparable to microchannel dimensions. Populations of nanospheres formed by the microfluidic PIN methods can be statistically grouped into two homogeneous subsets of 0.01% PMMA and less than 0.01% PMMA. Additionally, the populations of nanospheres formed at 0.01% PMMA vary significantly between the two microfluidic methods (p < 0.01), while at lower concentrations they are negligibly different (not statistically significant). The variation seen between the two microfluidic methods at higher concentrations evidence the extreme sensitivity to manufacturing conditions that have substantially complicated previous PIN-based processes. The mean volume decrease is negligible in both microfluidic testing conditions further evidencing a nonlinear relationship between polymer concentration and resultant particle size. On the whole, the flow pinching method tended to produce smaller spheres at the same polymer concentrations. While the flow rate of the solvent phase is slower in the flow pinching case than the nonflow pinching case, pinching the flow causes thinning of the solvent phase stream effectively increasing the interfacial area to volume ratio and allowing for more rapid diffusion-driven exchange between the solvent and nonsolvent. The increase in transport kinetics from the nonflow pinching to the flow pinching case would tend to accelerate polymer chain collapse leading to less trapped solvent, which may account for the observed size differences. The nanosphere populations at the concentrations below 0.01% produced by both microfluidic methods under the same conditions were very similar. Since the flow pinching method introduces the polymer solution into the nonsolvent rapidly it appears that the kinetics of nanosphere formation is faster still as the mean diameter and distribution of particles is similar in both microfluidic setups. Results indicate that the conditions for formation at 0.001 and 0.0001 wt % PMMA in methylene chloride are similar as evidenced by the similarities of resultant nanosphere populations formed by both microfluidic methods. Between 0.001 and 0.01 wt % there is a significant increase in mean nanosphere diameter.
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Figure 8. (a) Nanosphere population size histograms produced by the glass capillary tube method compared with both microchip production methods formed using the same polymer concentration. (b) Mean diameter of nanospheres produced by the glass capillary tube method plotted as a function of flow rate. A linear best fit line is shown. ** ) Statistical difference between 1 and 10 nL/s flow rates, p < 0.01; † ) 100 nL/s is statistically different from 1 and 10 nL/s flow rates, p < 0.001.
The overlap concentration (c*) for PMMA in a good solvent was calculated to be c* ≈ 1.2 weight per volume percent. Additionally the viscosities of all of the polymer solutions used was negligibly different from pure methylene chloride indicating that all of the studies were performed in perfectly dilute solution conditions.
Since only the highest concentration formed nanospheres offchip perhaps the polymer-solvent and nonsolvent phases are miscible at the lower polymer concentrations when mixed rapidly. At low flow rates in the laminar regime the solvent phase is introduced slowly and so the volume of the solvent phase is
Continuous Flow Nanosphere Formation
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Table 1. Nanosphere Population Characteristics Including Mean Diameter, the Diameter of a Sphere that 90% of the Population is Larger than (90% > x), the Diameter of a Sphere that 90% of the Population is Smaller than (90% < x), and the Percentage of the Population of Nanospheres that are within the Size Range Ideal for Non-phagocytotic Cellular Uptake (200-1000 nm in Diameter)
method
polyer concentration (wt %)
solvent phase flow rate (nL/s)
mean diameter (nm)
90% > x (nm)
90% < x (nm)
200-1000 nm (%)
off chip flow pinching microchip flow pinching microchip flow pinching microchip no flow pinching microchip no flow pinching microchip no flow pinching microchip glass capillary tube glass capillary tube glass capillary tube
0.01 0.0001 0.001 0.01 0.0001 0.001 0.01 0.001 0.001 0.001
uncontrolled/high 0.0932 0.0932 0.0932 0.3141 0.3141 0.3141 1 10 100
660 517 533 731 525 529 846 508 431 334
419 417 370 415 365 328 353 243 234 207
1047 603 695 987 652 791 1628 761 647 526
88.0 100.0 100.0 90.9 97.6 98.6 69.9 91.8 96.7 89.2
greatly reduced relative to that of the nonsolvent phase at any given time. In this case the nonsolvent to solvent ratio is effectively very high throughout the phase inversion process producing smaller and more uniform spheres due to the controlled mixing conditions. Capillary Tube Nanosphere Production. To further investigate the effect of solvent phase flow rate, the flow was varied and polymer concentration was held constant in the glass capillary tube experiments. The results show a statistically significant (p < 0.01) decreasing trend in mean nanosphere diameter with increasing flow rate (Figure 8). While the bulk flow rate was increased when moving from a flow-pinching to a nonflow pinching setup in the microfluidic case, the effective interfacial area involved in diffusion decreased indicating that both area for solvent nonsolvent exchange and flow rate can be tuned to control the resultant size distribution of the nanosphere population. The population of nanospheres produced by the glass capillary tube method at 100 nL/s flow rate produced a population of nanospheres with the greatest percentage in the 200-500 nm diameter range, 78.5%, of all the methods tested. Moreover, the population of nanospheres formed by the glass capillary tube method flowing the solvent phase at 1 nL/s has no statistically significant different (P < 0.01) from that formed by the microfluidic methods. For both microfluidic methods decreasing polymer concentration from 0.01 to 0.0001 wt % demonstrated an increase in size monodispersity with decreasing polymer concentration. However, with the decrease in concentration comes an increase in the required amount of organic solvent to yield the same final weight of product. If the cost of the therapeutic agent outweighs the cost of the organic solvent, as it typically does, then the increase in production within the size range of interest for improving bioavailability greatly outweighs the cost. Based on above results, we hypothesize two mechanisms (1) spinoidal decomposition and (2) nucleation and growth could explain the PIN phenomenon. In the spinoidal decomposition process polymer molecules instantaneously collapse upon phase inversion (exchange of solvent and nonsolvent) to form nanoparticles. In the nucleation and growth process, nucleation sites are created by inhomogeneous mixture sites around which polymer chains collapse to form polymer-rich and polymer-poor regions that after phase inversion yield solid polymer nanospheres. To test above hypothesizes of nanoparticle formation we varied the polymer phase flow rate within the laminar regime given that the mixing time is independent of flow rate. The experiments show a trend toward decreasing size with increasing flow rate indicating time-course-dependent nucleation and growth mechanism of formation for the resultant nanosphere population within the range of conditions tested. If the mechanism were instan-
taneous, flow rate is not expected to affect the resultant population. Additionally the formation of polymer network structures at higher concentrations morphologically suggests crowded nucleation sites that join as polymer chain collapse occurs as water freezes around nucleation sites to form snow flakes. Two competing theories have been used to explain PIN: nucleation and growth and spinoidal decomposition. The linear dependence of mean diameter on flow rate supports the time course dependent mechanism, nucleation and growth, as does the formation of the polymer network within the microchannels observed at higher polymer concentrations.
Conclusions Polymer nanospheres formulated in organic solvents were produced for the first time on a microfluidic platform by the PIN method. All but two of the described experimental conditions produced populations of nanospheres with at least 90% in the desired size range for uptake by nonphagocytitic cells, 200-1000 nm as shown in Table 1. Microfluidic investigations show a nonlinear dependence of population mean diameter on polymer concentration. In the lower range of concentrations tested particle size appeared to be insensitive to concentration evidencing an optimal concentration that will minimize solvent usage and maximize throughput for nanospheres in the desired size range. Whereas mean nanosphere diameter varied linearly with flow rate in the glass capillary tube studies within the range of flow rates from 1-100 nL/s, making solvent phase flow rate a very useful control parameter for further tuning resultant size distribution given an optimal concentration within the range of flow rates examined. Moreover, the experimental setups have and will continue to shed light on the mechanism of nanosphere production by PIN. In conclusion, PIN performed in laminar flow conditions yields polymer nanospheres of the optimal size for oral drug delivery to absorptive GI epithelial cells.1,27 Glass chip microfluidics provides a platform in which conditions for polymer nanosphere formation can be studied quickly and inexpensively. The glass microfluidic platform will prove useful for rapidly investigating the experimental parameters necessary for producing polymer nanospheres containing therapeutic agents. Production scale-up is easily achieved both in series (using larger channels) and in parallel (using many chips or glass capillary tube setups at once) allowing for continuous-flow production. The population of nanospheres produced by the glass capillary tube method using 0.001 wt % PMMA at a flow rate of 1 nL/s is statistically similar (27) Norris, D. A.; Puri, N.; Sinko, P. J. The effect of physical barriers and properties on the oral absorption of particulates. AdV. Drug DeliVery ReV. 1998, 34 (2-3), 135-154.
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(P < 0.05) to that produced by the flow pinching microchip method at the same concentration with a solvent flow rate of 0.0932 nL/s flow rate indicating a link between the two platforms that could be used to enable testing production parameters in small batches and then rapidly scaling production. From a mechanistic standpoint that flow pinching produces a nanospheres population more similar to the glass capillary tube method than the no flow pinching setup indicates that the availability of nonsolvent influences the resultant size distribution, which also supports a nucleation and growth mechanism of formation. Additionally, the glass microfluidics platform and simple chip design will allow for integration of other applications (e.g., dissolution profiling) on the same chip yielding laboratory-ona-chip technologies that will greatly accelerate data acquisition regarding nanosphere formation by phase inversion. Data obtained from varying flow rate in the glass capillary tube experiments suggests that PIN formation of nanospheres under the conditions
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investigated is mediated by nucleation and growth. As the mechanisms and kinetics of cellular nanosphere uptake are elucidated, controlled transport PIN will offer a technique for drug delivery scientists to improve cellular uptake of the increasing number of hydrophobic and biologic new chemical entities. Controlling the introduction of the solvent phase into the nonsolvent improves the reproducibility of manufacturing conditions. Through introducing microfluidic-controlled transport conditions we aim to determine the effect of three flow configurations of polymer and non solvent on the mean particle diameter and to explore mechanistic hypotheses of PIN polymer nanosphere formation. Acknowledgement The authors gratefully acknowledge that this project was supported by Brown University and thank Cartney Smith for his contributions to the project. LA8009332
