Langmuir 2008, 24, 9775-9783
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Poly(D,L-lactide-co-glycolide) Nanoparticle Agglomerates as Carriers in Dry Powder Aerosol Formulation of Proteins Laura J. Peek,†,‡ Lydia Roberts,‡ and Cory Berkland*,†,‡ Department of Chemical and Petroleum Engineering, and Department of Pharmaceutical Chemistry, UniVersity of Kansas, Lawrence, Kansas 66047 ReceiVed April 16, 2008. ReVised Manuscript ReceiVed June 10, 2008 A dry powder aerosol drug delivery system was designed with both nano- and microstructure to maximize the protein loading via surface adsorption and to facilitate delivery to the deep lung, respectively. Ovalbumin was employed as a model protein to adsorb to and controllably flocculate DOTAP-coated PLG nanoparticles into “nanoclusters” possessing low density microstructure. The mechanism of nanoparticle flocculation was probed by evaluating the effects of ionic strength, shear force, and protein concentration on the geometric and aerodynamic diameters of the nanoclusters as well as the protein adsorption efficiency. Salt ions were found to compete with ovalbumin adsorption to nanoparticles and facilitate flocculation; therefore, formulation of nanoclusters for inhaled drug delivery may require the lowest possible ionic strength to maximize protein adsorption. Additional factors, such as shear force and total protein-particle concentration can be altered to optimize nanocluster size, suggesting the possibility of regional lung delivery. Immediate release of ovalbumin was observed, and native protein structure upon release was confirmed by circular dichroism and fluorescence spectroscopy studies. Controlled flocculation of nanoparticles may provide a useful alternative to spray drying when formulating dry powders for pulmonary or nasal administration of protein therapeutics or antigens.
1. Introduction Inhaled aerosols are an effective means to treat diseases infecting the lung, including asthma and cystic fibrosis, which require treatment locally in the bronchial or alveolar region.1,2 Alternatively, the large surface area of the pulmonary system (∼140 m2), attributable to the alveolar epithelium in the deep lung region, and ready access to the circulation offer the potential for noninvasive systemic delivery of macromolecular therapeutics including peptides, proteins, DNA, etc.3–8 Efficiently administering aerosols to the deep lung remains an encumbrance when locally treating lung tissue or when utilizing the lung as a gateway for systemic drug delivery.9 Deposition of inhaled particles in the deep lung requires significant control of the particle size, a major deterrent to utilizing the lung for drug delivery.9–11 Particles possessing an aerodynamic diameter greater than 15 µm are subject to deposition in the mouth and throat, while submicron particles are commonly exhaled.12–14 Studies have consistently verified that ∼1-3 µm * Corresponding author. Address: 2030 Becker Drive, Lawrence, KS 66047;Telephone:785-864-1455;Fax:785-864-1454;E-mail:
[email protected]. † Department of Chemical and Petroleum Engineering. ‡ Department of Pharmaceutical Chemistry.
(1) Niven, R. W. Crit. ReV. Ther. Drug Carrier Syst. 1995, 12, 151–231. (2) Reed, C. E.; Offord, K. P.; Nelson, H. S.; Li, J. T.; Tinkelman, D. G. J. Allergy Clin. Immunol. 1998, 101, 14–23. (3) Bivas-Benita, M.; Romeijn, S.; Junginger, H. E.; Borchard, G. Eur. J. Pharm. Biopharm. 2004, 58, 1–6. (4) Happel, K. I.; Lockhart, E. A.; Mason, C. M.; Porretta, E.; Keoshkerian, E.; Odden, A. R.; Nelson, S.; Ramsay, A. J. Infect. Immun. 2005, 73, 5782–5788. (5) Patton, J. S. CHEMTECH 1997, 27, 34–38. (6) Patton, J. S. AdV. Drug DeliVery ReV. 2000, 42, 239–248. (7) Patton, J. S.; Platz, R. M. AdV Drug DeliVery ReV. 1992, 8, 179–196. (8) Scott, E. S.; Wiseman, J. W.; Evans, M. J.; Colledge, W. H. J. Gene Med. 2001, 3, 125–134. (9) Edwards, D. A.; Hanes, J.; Caponetti, G.; Hrkach, J.; Ben-Jebria, A.; Eskew, M. L.; Mintzes, J.; Deaver, D.; Lotan, N.; Langer, R. Science 1997, 276, 1868– 1872. (10) Newman, S. P. Curr. Opin. Pulm. Med. 2003, 9 Suppl 1, S17–20. (11) Newman, S. P.; Hirst, P. H.; Pitcairn, G. R. Curr. Opin. Pulm. Med. 2001, 7 Suppl 1, S12–14. (12) Musante, C. J.; Schroeter, J. D.; Rosati, J. A.; Crowder, T. M.; Hickey, A. J.; Martonen, T. B. J. Pharm. Sci. 2002, 91, 1590–1600.
solid particles deposit most efficiently in the deep lung, that is the bronchiole and alveolar region.12–14 In light of this, low density particles are being developed in several laboratories as a means to deliver drugs to the deep lung.9,15–17 These particles possess large geometric diameters (dp), but because of their low mass densities (F) exhibit much smaller aerodynamic diameters (daero) as described by eq 1,15
daero ) dp[(F/Fref)/γ]0.5
(1)
where Fref is a reference mass density (e.g., 1 g/cm3) and γ is a dynamic shape factor (typically 1 for a sphere). In addition to improvements in aerosol microstructure, other studies have focused on developing nanoparticles as drug delivery vehicles to the pulmonary system.18,19 Nanoparticles may not be desirable as aerosols since submicron particles are often exhaled following administration.12–14 Particles of this size also suffer from a propensity to agglomerate uncontrollably,20,21 thus reducing their utility as aerosols. An advantage that nanoparticles offer to pulmonary drug delivery is the increased surface area (per total particle mass) available for drug adsorption.22 Moreover, macromolecules adsorbed to the surface of nanoparticles may (13) Pritchard, J. N. J. Aerosol Med. 2001, 14, S19–26. (14) Lipworth, B. J. Respir. Med. 2000, 94, S13–16. (15) Edwards, D. A.; Ben-Jebria, A.; Langer, R. J. Appl. Physiol. 1998, 85, 379–385. (16) Vanbever, R.; Mintzes, J. D.; Wang, J.; Nice, J.; Chen, D.; Batycky, R.; Langer, R.; Edwards, D. A. Pharm. Res. 1999, 16, 1735–1742. (17) Fu, J.; Fiegel, J.; Krauland, E.; Hanes, J. Biomaterials 2002, 23, 4425– 4433. (18) John, A. E.; Lukacs, N. W.; Berlin, A. A.; Palecanda, A.; Bargatze, R. F.; Stoolman, L. M.; Nagy, J. O. J. FASEB 2003, 17, 2296–2298. (19) Kumar, M.; Kong, X.; Behera, A. K.; Hellermann, G. R.; Lockey, R. F.; Mohapatra, S. S. Genet. Vaccines Ther. 2003, 1, 3. (20) Bozdag, S.; Dillen, K.; Vandervoort, J.; Ludwig, A. J. Pharm. Pharmacol. 2005, 57, 699–707. (21) Quintanar-Guerrero, D.; Ganem-Quintanar, A.; Allemann, E.; Fessi, H.; Doelker, E. J. Microencapsul. 1998, 15, 107–119. (22) Wendorf, J.; Singh, M.; Chesko, J.; Kazzaz, J.; Soewanan, E.; Ugozzoli, M.; O’Hagan, D. J. Pharm. Sci. 2006, 95, 2738–2750.
10.1021/la8012014 CCC: $40.75 2008 American Chemical Society Published on Web 08/05/2008
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offer the advantage of improved stability and activity as compared to encapsulated proteins.23 Poly(D,L-lactide-co-glycolide) (PLG) is an attractive polymer for fabricating nano- and microparticles for drug delivery because of its biodegradability and biocompatibility;24 however, despite its approved use in parenteral formulations, the use of PLG in inhaled drug formulations remains to be accepted. Concerns surrounding the slow degradation rate of PLG and the potential for accumulation of the polymer in the lungs are obstacles in the development of such pulmonary delivery vehicles that remain to be evaluated;25–27 however, recent work by Kissell and others demonstrate the safety of PLG nanoparticles compared to nondegradable nanoparticles (polystyrene) in rats.27 We have designed an inhalable dry powder formulation possessing both nano- and microstructure to increase adsorption of a macromolecular drug and improve delivery to the deep lung, respectively. Surface charge interactions were exploited for controlled flocculation of nanoparticles using ovalbumin as a model protein drug, and various properties of the nanoparticles were evaluated for their effect on protein adsorption and flocculate properties. Factors such as surface charge of the nanoparticles, ionic strength, shear force, protein-particle ratio, and total protein-particle concentration were found to influence the formation, size, and density of the flocculated nanoparticles, or “nanoclusters.” Ovalbumin was immediately released from the lyophilized nanoclusters into 10 mM sodium phosphate buffer (pH 7.0) containing 150 mM NaCl (PBS) and maintained native protein structure as determined by fluorescence and circular dichroism (CD) measurements. This formulation approach leverages agglomeration of nanoparticles in solution as protein carriers and may serve as an alternative to spray drying when formulating dry powder aerosols. Ultimately, we believe that the nanocluster delivery system may enhance the transport of macromolecules to the deep lung for both local and systemic delivery with the potential for delivering synergistic, combination therapeutics.
2. Materials and Methods 2.1. Materials. PLG (50:50, 0.41 dl/g) was purchased from Lactel (Pelham, AL). Poly(vinyl alcohol) (88 mol% hydrolyzed; PVA) was purchased from Polysciences, Inc. (Warrington, PA). 1,2Dioleoyl-3-trimethylammoniopropane, chloride salt (DOTAP) and 1,2-dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) were purchased from Avanti Polar Lipids, Inc. (Alabaster, AL). Albumin from chicken egg white (ovalbumin, grade V, minimum 98%) was purchased from Sigma (St. Louis, MO). All other reagents were purchased in high purity from Fisher Scientific (Pittsburgh, PA) and used without further purification. Aqueous solutions of each lipid were prepared by first drying and then redissolving the lipid in water. Briefly, lipid in chloroform was added to a volumetric flask. A stream of nitrogen was used to dry the lipid in the bottom of the flask while gradually rotating the flask to create a film. Distilled, deionized water (ddH2O) was added to the flask containing the dry lipid film, and the flask was swirled and sonicated to mix and dissolve the lipid. 2.2. Fabrication of Charged Nanoparticles. A modified emulsion solvent evaporation method28,29 was employed to create cationic, (23) Duncan, G.; Jess, T. J.; Mohamed, F.; Price, N. C.; Kelly, S. M.; van der Walle, C. F. J. Controlled Release 2005, 110, 34–48. (24) Jain, R. A. Biomaterials 2000, 21, 2475–2490. (25) Pandey, R.; Khuller, G. K. J. Antimicrob. Chemother. 2005, 55, 430–435. (26) Dunne, M.; Corrigan, I.; Ramtoola, Z. Biomaterials 2000, 21, 1659– 1668. (27) Dailey, L. A.; Jekel, N.; Fink, L.; Gessler, T.; Schmehl, T.; Wittmar, M.; Kissel, T.; Seeger, W. Toxicol. Appl. Pharmacol. 2006, 215, 100–108. (28) Kazzaz, J.; Neidleman, J.; Singh, M.; Ott, G.; O’Hagan, D. T. J. Controlled Release 2000, 67, 347–356. (29) Mainardes, R. M.; Evangelista, R. C. J. Microencapsul. 2005, 22, 13–24.
Peek et al. anionic and neutral nanoparticles. Briefly, nanoparticles were fabricated by sonicating (50% amplitude) or homogenizing (30 000 rpm) 3 mL of 1.67% (w/v) PLG dissolved in an acetone/methanol (5/1) solvent mixture into 25 mL aqueous solution containing one of the following: DOTAP (cationic), dioctyl sulfosuccinate (DSS) (anionic), cetyltrimethylammonium bromide (CTAB) (cationic), or PVA (neutral). DOTAP and CTAB nanoparticles were prepared using sonication, while all others were prepared using homogenization. The suspension was stirred for 4 h to evaporate the solvent. The precipitated nanoparticles were then washed with ddH2O and sonicated in a water bath to break up any loosely associated agglomerates. The nanosuspension was then filtered through a KimWipe laboratory tissue to remove large, particulate matter (i.e., agglomerates, film-like residue, etc.) and characterized. 2.3. Characterization of Nanoparticles. The geometric diameters and polydispersities of the nanosuspensions were evaluated using dynamic light scattering (Brookhaven Instruments Zeta Potential Analyzer, Holtsville, NY). Nanoparticles were diluted in ddH2O (∼100×), and three 1 min measurements were obtained at 25 °C for each nanosuspension. The same instrument was employed to quantify the zeta potential, an indicator of surface charge, of the nanoparticles in 1 mM sodium nitrate solution. Three runs of 15 cycles were acquired, and the mean zeta potential (ζ) was recorded. 2.4. Adsorption of Ovalbumin to Charged Nanoparticles. Ovalbumin was employed as a model macromolecular drug for these studies. Working solutions of ovalbumin in PBS were prepared, and the exact concentration of each was determined by UV absorbance spectroscopy (Agilent 8453) using the absorbance at 280 nm and an extinction coefficient of 0.696 mL/mg · cm. Aliquots (100 µL) of ovalbumin working solutions were added to a constant mass of cationic, anionic, and neutral nanoparticles (1 mL) in 2 mL microcentrifuge tubes (final salt concentration was 14 mM). Samples were prepared in triplicate and tumbled end-over-end for 1 h at 4 °C to facilitate adsorption. Centrifugation (14 000g) was used to pellet the particles, and the supernatant was assayed for unbound ovalbumin using UV absorbance spectroscopy. Total surface area was estimated using the following process:30 (1) the mean diameter measured by dynamic light scattering was used to calculate the surface area of a single nanoparticle, (2) the mass of a single nanoparticle was estimated using the volume of a single nanoparticle and the absolute density of PLGA, measured previously in our laboratory using a helium pycnometer,31 (3) the concentration of nanoparticles (determined by lyophilizing a known volume of the nanosuspension in a preweighed tube and then weighing the mass of particles postlyophilization) and the mass of a single particle was used to calculate the total number of particles, and (4) the surface area of a single nanoparticle was multiplied by the total number of particles to arrive at the total surface area. Although the calculations do not provide an absolute value for total adsorptive surface area, we do believe that this approach yields a close approximation. 2.5. Flocculation of DOTAP Nanoparticles with Ovalbumin. 2.5.1. Effects of OValbumin Concentration, Ionic Strength, and Shear Force during Tumbling on Flocculation. The mechanism of flocculation of the DOTAP nanoparticles in the presence of ovalbumin was probed by comparing flocculates prepared in ddH2O and PBS to evaluate effects of ionic strength. Additionally, the effect of headspace during tumbling (i.e., shear force) was evaluated by comparing the size of the resulting flocculates prepared in 2 mL microcentrifuge tubes and 15 mL centrifuge tubes. To each tube, ovalbumin working solution (408 µL) in either ddH2O or PBS was added to 400 µL of DOTAP-coated nanoparticles (1.16 mg) to yield a weight ratio of ovalbumin to nanoparticles on the order of approximately 0, 0.01, 0.10, 0.30, and 0.60. The concentrations of ovalbumin working solutions were 0, 0.028, 0.28, 0.85, 1.7 mg/mL, respectively. Blanks (without ovalbumin) were prepared to evaluate the role of the buffer salts in flocculation. The final salt concentration (30) Zhang, N.; Chittasupho, C.; Duangrat, C.; Siahaan, T. J.; Berkland, C. Bioconjugate Chem. 2008, 19, 145–152. (31) Arnold, M. M.; Gorman, E. M.; Schieber, L. J.; Munson, E. J.; Berkland, C. J. Controlled Release 2007, 121, 100–109.
PLG Nanoparticle Agglomerates as Carriers in all samples prepared using PBS was ∼81 mM. The samples were tumbled end-over-end at 4 °C for 1 h, and the geometric diameters of the resulting flocculates were measured using a Multisizer 3 Coulter Counter (Beckman Coulter, Fullerton, CA). Additionally, the effect of protein adsorption on the surface charge was evaluated using a zeta potential analyzer. Forty microliters of ovalbumin-adsorbed nanoparticles were added to 2 mL of 1 mM sodium nitrate solution. The zeta potential was quantified as described above. Furthermore, the remaining sample was centrifuged to pellet the particles, and the supernatant was assayed for unbound protein concentration using UV absorbance spectroscopy. 2.5.2. Effects of Total Protein-Particle Concentration on Flocculation. A weight ratio of 0.66 ovalbumin to nanoparticles and a total mass of 1.74 mg of ovalbumin and nanoparticles were maintained for all samples. Aliquots of the DOTAP-coated nanoparticle suspension (400 µL) were added to 15 mL centrifuge tubes and diluted by adding 500 µL, 2, 5, or 10 mL of PBS or ddH2O. A 408 µL aliquot of the appropriate 2.81 mg/mL ovalbumin working solution (PBS or ddH2O) was then added to each tube to yield a final total concentration of 1.3, 0.62, 0.33, and 0.16 mg/mL, respectively. The samples were treated as described above to facilitate flocculation and analyzed using a Multisizer 3 Coulter Counter to measure the geometric diameters of the flocculates. 2.6. Dry Powder Nanoclusters. Flocculates (808 µL) were prepared in triplicate in 2 mL microcentrifuge tubes containing a weight ratio of ovalbumin to nanoparticles of ∼0.01, 0.10, 0.30, and 0.60 in either ddH2O or PBS as described above. The final ionic strength of the nanosuspensions prepared using PBS was ∼81 mM. To maximize sample recovery, the flocculates from each of the three 2 mL tubes were combined into a 15 mL centrifuge tube, frozen at -80 °C, and lyophilized using a Labconco benchtop lyophilizer (Kansas City, MO). 2.6.1. Characterization of Nanocluster Particle Morphology. The size and morphology of the dry powder nanoclusters were evaluated using a LEO 1550 field emission scanning electron microscope (SEM) with secondary electron detection. The flocculates were coated on a platform and sputtered with gold prior to imaging at 10 000× magnification. 2.6.2. Characterization of Nanocluster Aerosol Properties. The aerodynamic diameters of the dry powder nanoclusters were evaluated using an Aerosizer LD (Amherst Process Instruments Inc.). Data were collected over ∼70 s using a 300 µm nozzle and low shear force. 2.6.3. Tap Test Method To Determine Bulk Powder Density. The bulk density of the dry powder was estimated using a microtap test approach. Briefly, dry powder nanoclusters were added to preweighed microcentrifuge tubes, and the tubes were weighed back to determine the mass of powder. The tubes were then tapped on the benchtop 30 times to compress the dry powder. The volume of the powder was approximated by comparing the compressed powder level to the level of a known volume of water. The density was calculated by dividing the mass of powder by the approximated volume. Samples were analyzed in triplicate in most cases, but because of a sample shortage, the powder at a protein-particle ratio of 0.28 prepared in ddH2O was only evaluated in duplicate. The bulk powder densities of the nanoclusters prepared in ddH2O at protein-particle ratios of 0.02 and 0.10 were not quantified because of lack of sample. 2.7. Release of Ovalbumin from Nanoclusters. Studies were conducted to evaluate the release kinetics of ovalbumin from the dry powder nanoclusters. Release was monitored in PBS and in an aqueous suspension of DPPC vesicles (13.25 mg/mL). Nanoclusters (78.1 and 35.6 mg) were suspended in 15 mL of PBS and 10 of mL DPPC, respectively. Aliquots (500 µL) were transferred to 2 mL microcentrifuge tubes and incubated at 37 °C while shaking at 175 rpm. Duplicate samples (time ) 0) were immediately centrifuged at 13 000g for 30 min to pellet the particles. The supernatant was assayed for protein concentration by UV absorbance spectroscopy (supernatant from DPPC samples were diluted 2× to minimize interference with absorbance near 280 nm). After removal of the supernatant, the remaining pellet was dissolved in 40 µL of dimethyl sulfoxide (DMSO), and the polymer was then precipitated by addition
Langmuir, Vol. 24, No. 17, 2008 9777 Table 1. Characteristics of Nanoparticles (N ) 3) coating material, concentration
sizea (nm)
zeta potential (ζ, mV)
DOTAP, 50 µM CTAB, 0.5% (w/v) PVA, 0.5% (w/v) DSS, 0.1% (w/v)
271.5 ( 9.1 296.2 ( 4.1 182.3 ( 2.1 313.0 ( 18.9
42.6 ( 3.2 23.0 ( 1.7 -2.6 ( 3.2 -36.4 ( 2.1
a
The polydispersity values were less than 0.16 in all cases.
of 350 µL of PBS. The samples were centrifuged at 13 000g for 10 min to pellet the polymer, and the supernatant was again assayed for unreleased protein concentration by UV absorbance spectroscopy. 2.7.1. Stability of Released OValbumin. After flocculate formation and prior to lyophilization, the flocculates were washed with ddH2O to remove excess, unbound ovalbumin. To wash the flocculates, the samples were centrifuged at 500 rpm for 5 min. The supernatant was discarded, and the flocculates were resuspended in ddH2O. This was repeated, and the washed flocculates were resuspended in ∼2 mL of ddH2O, frozen at -80 °C, and lyophilized. Since immediate release of ovalbumin from the nanoclusters was observed (see Results), the lyophilized nanoclusters were resuspended in 1 mL of PBS to release ovalbumin and centrifuged at 14 000g for 10 min to pellet the particles. The supernatant was isolated and analyzed using UV absorbance, fluorescence (PTI QuantMaster, Lawrenceville, NJ), and circular dichroism (CD; Jasco J-810, Easton, MD) spectroscopy to evaluate the conformational stability of the released ovalbumin. UV absorbance was employed to determine the concentration of ovalbumin in the samples. Fluorescence studies utilized the tryptophan (Trp) residues of ovalbumin as an intrinsic fluorescence probe sensitive to the polarity of their environment to determine whether the tertiary structure of ovalbumin was altered during flocculation. Fluorescence spectra of the released ovalbumin and a control containing ovalbumin in PBS were collected from 305-400 nm using an excitation wavelength of 295 nm (>95% Trp emission). CD was employed to evaluate whether the secondary structure of ovalbumin was altered during flocculation. CD spectra of the sample and control were acquired in three accumulations from 260-195 nm with a scanning speed of 50 nm/min and 1.0 nm resolution.
3. Results 3.1. Characterization of Charged Nanoparticles. The sizes and surface charges of PLG nanoparticles were modified using four different charged coating materials (Table 1). Particles of approximately 200-300 nm were created with relatively low polydispersity values (
