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Quick Synthesis of Lipid-Polymer Hybrid Nanoparticles with Low Polydispersity Using a Single-Step Sonication Method Ronnie H. Fang, Santosh Aryal, Che-Ming Jack Hu, and Liangfang Zhang* Department of Nanoengineering and Moores Cancer Center, University of California-San Diego, La Jolla, California 92093, United States Received September 6, 2010. Revised Manuscript Received September 26, 2010 Lipid-polymer hybrid nanoparticle, consisting of a hydrophobic polymeric core and a lipid monolayer shell, represents a new and promising drug delivery platform that has shown controllable particle size and surface functionality, high drug loading yield, sustained drug release profile, and excellent in vitro and in vivo stability. These lipid monolayer-coated polymeric nanoparticles are typically fabricated through a modified nanoprecipitation method, which involves sample heating, vortexing, and solvent evaporation. Herein we report a new and fast method to synthesize lipid-polymer hybrid nanoparticles with controllable and nearly uniform particle size. Using a bath sonication approach, we demonstrate that the whole hybrid nanoparticle synthesis process can be completed in about 5 min compared with a few hours for previous synthesis approaches. The size and polydispersity of the resulting nanoparticles can be readily controlled by tuning the relative concentrations of individual building components. Colloidal stability tests of the synthesized hybrid nanoparticles in PBS buffer and serum show no signs of aggregation over a period of 5 days. The present method improves the production rate of the hybrid nanoparticles by near 20-fold while not compromising the physicochemical properties of the particles. This work may facilitate the bench-to-bedside translation of lipid-polymer hybrid nanoparticles as a robust drug nanocarrier by allowing for fabricating a large amount of these nanoparticles at high production rate.
Introduction The advent of nanoparticle technology has brought new hope for the treatment of various notorious diseases such as cancers, cardiovascular diseases, and bacterial infections.1-5 As a delivery vehicle, nanoparticles with a size range of 10-150 nm have shown great advantages compared with the conventional therapeutics. These advantageous properties include improving the solubility of poorly water-soluble drugs, prolonging the circulation lifetime of drugs, releasing drugs at a sustained rate and thus lowering the frequency of administration, delivering drugs in a targeted manner to minimize systemic side effects, and delivering multiple types of drugs simultaneously for combination therapy.6-12 As a result, many nanoparticle platforms have been designed for drug delivery applications. Over two dozen nanoparticle-based therapeutic
products have been approved for clinical use, and numerous other ensuing products are in clinical trials.13-15 Among these clinical or preclinical products, liposomes9,16 and biodegradable polymeric nanoparticles17,18 represent two dominant classes of drug nanocarriers. Recently, efforts have been made to combine the positive attributes of both liposomes and polymeric nanoparticles into a single delivery system, named lipid-polymer hybrid nanoparticles.19-25 This type of nanoparticle is typically comprised of three distinct functional components: a hydrophobic polymeric core to encapsulate poorly watersoluble drugs with high loading yields; a lipid layer surrounding the core to provide a highly biocompatible shell and to promote drug retention inside the polymeric core; and a hydrophilic polymer stealth layer outside the lipid shell to enhance nanoparticle stability and systemic circulation lifetime. Depending on how these hybrid nanoparticles are prepared, the lipid shell can be a
*Corresponding author: e-mail
[email protected], Tel 858-246-0999, Fax 858-534-9553. (1) Brannon-Peppas, L.; Blanchette, J. O. Adv. Drug Delivery Rev. 2004, 56, 1649–1659. (2) Farokhzad, O. C.; Langer, R. Adv. Drug Delivery Rev. 2006, 58, 1456–1459. (3) Kawasaki, E. S.; Player, A. Nanomedicine 2005, 1, 101–109. (4) Peer, D.; Karp, J. M.; Hong, S.; Farokhzad, O. C.; Margalit, R.; Langer, R. Nature Nanotechnol. 2007, 2, 751–760. (5) Zhang, L.; Pornpattananangku, D.; Hu, C. M.; Huang, C. M. Curr. Med. Chem. 2010, 17, 585–594. (6) Cuong, N. V.; Hsieh, M. F. Curr. Drug Metab. 2009, 10, 842–850. (7) Hu, C. M.; Zhang, L. Curr. Drug Metab. 2009, 10, 836–841. (8) Tong, R.; Cheng, J. J. Polym. Rev. 2007, 47, 345–381. (9) Torchilin, V. P. Nat. Rev. Drug Discovery 2005, 4, 145–160. (10) Hu, C.-M.; Aryal, S.; Zhang, L. Ther. Delivery 2010, 1, 323–334. (11) Bouladjine, A.; Al-Kattan, A.; Dufour, P.; Drouet, C. Langmuir 2009, 25, 12256–12265. (12) Wang, Y.; Han, P.; Xu, H.; Wang, Z.; Zhang, X.; Kabanov, A. V. Langmuir 2010, 26, 709–715. (13) Davis, M. E.; Chen, Z. G.; Shin, D. M. Nat. Rev. Drug Discovery 2008, 7, 771–782. (14) Zhang, L.; Gu, F. X.; Chan, J. M.; Wang, A. Z.; Langer, R. S.; Farokhzad, O. C. Clin. Pharmacol. Ther. 2008, 83, 761–769. (15) Wagner, V.; Dullaart, A.; Bock, A.-K.; Zweck, A. Nature Biotechnol. 2006, 24, 1211–1217.
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(16) Abra, R. M.; Bankert, R. B.; Chen, F.; Egilmez, N. K.; Huang, K.; Saville, R.; Slater, J. L.; Sugano, M.; Yokota, S. J. J. Liposome Res. 2002, 12, 1–3. (17) Cheng, J.; Teply, B. A.; Sherifi, I.; Sung, J.; Luther, G.; Gu, F. X.; LevyNissenbaum, E.; Radovic-Moreno, A. F.; Langer, R.; Farokhzad, O. C. Biomaterials 2007, 28, 869–876. (18) Soppimath, K. S.; Aminabhavi, T. M.; Kulkarni, A. R.; Rudzinski, W. E. J. Controlled Release 2001, 70, 1–20. (19) Chan, J. M.; Zhang, L.; Tong, R.; Ghosh, D.; Gao, W. W.; Liao, G.; Yuet, K. P.; Gray, D.; Rhee, J. W.; Cheng, J. J.; Golomb, G.; Libby, P.; Langer, R.; Farokhzad, O. C. Proc. Natl. Acad. Soc. U.S.A. 2010, 107, 2213–2218. (20) Chan, J. M.; Zhang, L.; Yuet, K. P.; Liao, G.; Rhee, J. W.; Langer, R.; Farokhzad, O. C. Biomaterials 2009, 30, 1627–1634. (21) De Miguel, I.; Imbertie, L.; Rieumajou, V.; Major, M.; Kravtzoff, R.; Betbeder, D. Pharm. Res. 2000, 17, 817–824. (22) Liu, Y. T.; Li, K.; Pan, J.; Liu, B.; Feng, S. S. Biomaterials 2010, 31, 330– 338. (23) Sengupta, S.; Eavarone, D.; Capila, I.; Zhao, G. L.; Watson, N.; Kiziltepe, T.; Sasisekharan, R. Nature 2005, 436, 568–572. (24) Thevenot, J.; Troutier, A. L.; David, L.; Delair, T.; Ladaviere, C. Biomacromolecules 2007, 8, 3651–3660. (25) Zhang, L.; Chan, J. M.; Gu, F. X.; Rhee, J. W.; Wang, A. Z.; RadovicMoreno, A. F.; Alexis, F.; Langer, R.; Farokhzad, O. C. ACS Nano 2008, 2, 1696– 1702.
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lipid monolayer,25,26 bilayer, or multilayer.23,24,27 These lipidpolymer hybrid nanoparticles have been demonstrated to include some unique advantages of both liposomes and polymeric nanoparticles while excluding some of their intrinsic limitations, thereby holding great promise as a delivery vehicle for various medical applications. For the lipid-polymer hybrid nanoparticles with a lipid monolayer shell, they are usually prepared through a single-step nanoprecipitation method28,29 that induces self-assembly of the lipids and polymers.25 Typically, free polymers and hydrophobic drugs are dissolved in a water-miscible organic solvent such as acetonitrile, while lipids and lipid-PEG conjugates are dispersed in an aqueous solution. The polymer solution is then added into the lipid aqueous solution dropwise. When the organic solvent diffuses into the aqueous solution, the polymers will precipitate into small nanoparticles. Spontaneously, the lipids and lipidPEG will self-assemble on the surface of these polymer nanoparticles through hydrophobic interactions to reduce the system’s free energy. The hydrophobic tail of lipids will stick to the hydrophobic polymer core, and the hydrophilic headgroup of lipids will extend into the external aqueous environment. The lipid-PEG conjugate will also participate in the self-assembly process with its lipid moiety inserting into the lipid monolayer and its PEG moiety facing outward from the lipid monolayer to form a stabilizing and stealth corona of the nanoparticles. In this approach, the aqueous solution containing lipid and lipid-PEG needs to be heated to about 65 °C to ensure proper dispersion of lipids before adding in the polymer solution. The mixture solutions are usually vortexed for a few minutes followed by stirring for up to 2 h in order to uniformly coat the polymeric core with a lipid shell and to evaporate off some of the organic solvent.25 More recently, the above nanoprecipitation approach has been applied within a microfluidic rapid mixing device, which was able to synthesize both homogeneous core-shell structured lipidpolymer hybrid nanoparticles and lipid-quantum dots nanoparticles.30 Herein, we report an alternative approach to synthesize these lipid-polymer hybrid nanoparticles using a fast sonication method in place of heating, vortexing, and solvent evaporation. By adding all building components at predefined concentrations into a “cocktail”, followed by a short period of bath sonication, we notice that the lipids and polymers self-assemble to form nearly uniform nanoparticles. The particle size and size distribution can be readily optimized by tuning the concentration ratios among lipids, lipid-PEG, and polymers. The resulting hybrid nanoparticles remain stable in PBS buffer and serum. This sonication approach can fabricate lipid-polymer hybrid nanoparticles with a production rate about 20-fold higher than the existing method without compromising the physicochemical properties of the nanoparticles.
Experimental Section Materials. Ester-terminated poly(DL-lactic-co-glycolic acid) (PLGA) (inherent viscosity = 0.82 dL/g) was obtained from LACTEL Absorbable Polymers (Pelham, AL). 1,2-Distearoyl-snglycero-3-phosphoethanolamine-N-carboxy(poly(ethylene glycol)) (26) Hu, C. M.; Kaushal, S.; Cao, H. S.; Aryal, S.; Sartor, M.; Esener, S.; Bouvet, M.; Zhang, L. Mol. Pharmaceutics 2010, 7, 914–920. (27) Mornet, S.; Lambert, O.; Duguet, E.; Brisson, A. Nano Lett. 2005, 5, 281–285. (28) Aubry, J.; Ganachaud, F.; Cohen Addad, J. P.; Cabane, B. Langmuir 2009, 25, 1970–1979. (29) Gindy, M. E.; Panagiotopoulos, A. Z.; Prud’homme, R. K. Langmuir 2008, 24, 83–90. (30) Valencia, P. M.; Basto, P. A.; Zhang, L.; Rhee, M.; Langer, R.; Farokhzad, O. C.; Karnik, R. ACS Nano 2010, 4, 1671–1679.
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2000 (DSPE-PEG) was obtained from Avanti Polar Lipids (Alabaster, AL). Refined lecithin was obtained from Alfa Aesar (Ward Hill, MA). Fetal bovine serum, PBS buffer, acetonitrile, and all other solvents were purchased from Sigma-Aldrich (St. Louis, MO). Hybrid Nanoparticle Synthesis. Stock solutions of DSPEPEG and lecithin were prepared separately at concentrations of 1 mg/mL in 4 wt % ethanol aqueous solution. Stock solutions of PLGA were prepared at a concentration of 2.5 mg/mL in acetonitrile. The desired amounts of DSPE-PEG and lecithin solutions were added into deionized water. Then the PLGA solution was carefully pipetted into the resulting aqueous solution. Finally, the amount of deionized water used was adjusted so that the final volume ratio of aqueous to organic solution was 10:1. For example, to make hybrid nanoparticles (containing 1 mg of PLGA polymer) that have a lipid-PEG to PLGA polymer weight ratio of 0.30 and have 33% of the lipid-PEG replaced with its mole equivalent in lecithin, the following volumes were used: 400 μL stock PLGA solution, 200 μL DSPE-PEG stock solution, 12.5 μL lecithin stock solution, and 3787.5 μL deionized water. After everything was added together, the resulting mixture solution was sonicated in a capped glass vial for 5 min using a Fisher Scientific (FS30D) bath sonicator at a frequency of 42 kHz and power of 100 W. Purification was done by washing the solution in PBS buffer (pH = 7.4, 1X) 3 times using a Millipore (Amicon Ultra) centrifuge filter with a molecular weight cutoff of 10 kDa. For subsequent characterization, the resulting nanoparticles were concentrated to 1 mg of PLGA polymer per 1 mL of particle solution, denoted as 1 mg/mL, again using the same Amicon filter. Nanoparticle Characterization. In order to determine the particle characteristics (z-average diameter and polydispersity) and to obtain an intensity vs size plot, measurements were taken by dynamic light scattering (DLS) using the Malvern Zetasizer (ZEN 3600). All measurements were conducted at a backscattering angle of 173° and a temperature of 23 °C. Three subruns were conducted per measurement, and the average values were taken. Scanning electron microscopy (SEM) was used to visualize the size, size distribution, and morphology of the nanoparticles. Particle solutions (1 mg/mL) were diluted 500, and 5 μL of the diluted solutions was dropped onto a silicon wafer to dry overnight. The wafer with dried sample on them was then coated with chromium and imaged by SEM. Nanoparticle Stability Tests. To examine the long-term stability of the nanoparticles, they were suspended in 1X PBS buffer (ionic strength = 150 mM) with a concentration of 1 mg/mL. DLS was used to measure the particle size and polydispersity index (PDI) every 24 h for a period of 5 days. Serum stability tests were conducted by suspending the nanoparticles in 50% fetal bovine serum (FBS) with a final nanoparticle concentration of 1 mg/mL. In order to do this, particles were washed in PBS using Amicon filters as previously described and concentrated to 2 mg/mL. An equal volume of FBS was then added. As 50% FBS interfered with the DLS measurement, an absorbance method was used to monitor the particle size change in the presence of FBS.31 Absorbance measurements were conducted using a TECAN multiplate reader (Infinite M200). Samples were incubated at 37 °C with light shaking every 7 min inside the machine. The absorbance at 560 nm was taken approximately every 45 min over a period of 24 h. Bare PLGA nanoparticles with different size were used to calibrate the absorbance method. PLGA-PEG nanoparticles and serum alone (without nanoparticles) served as negative controls, while bare PLGA nanoparticles served as a positive control.
Results and Discussion Using PLGA, lecithin, and DSPE-PEG as representative hydrophobic polymer, lipid, and lipid-PEG conjugate, respectively, (31) Popielarski, S. R.; Pun, S. H.; Davis, M. E. Bioconjugate Chem. 2005, 16, 1063–1070.
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Figure 1. Synthesis and characterization of lipid-polymer hybrid nanoparticles. (A) Schematic illustration of the single-step sonication method to synthesize lipid-PLGA hybrid nanoparticles. At left, all of the components including PLGA polymer, lecithin lipid, and DSPE-PEG conjugates are mixed together as a “cocktail”. After 5 min of sonication, these building components self-assemble into core-shell structured nanoparticles. (B) Hydrodynamic size (diameter, nm) of the resulting hybrid nanoparticles measured by dynamic light scattering (DLS). (C) Representative scanning electron microscopy (SEM) images of the hybrid nanoparticles.
Figure 1A illustrates the process of preparing lipid-PLGA hybrid nanoparticles using a quick sonication method. The PLGA solution was directly added into the aqueous solution containing lecithin and DSPE-PEG with proper molar ratios and concentrations at room temperature. The final aqueous to organic solution was adjusted to a volume ratio of 10:1 by adding extra deionized water. The resulting mixture solution was then sonicated for 5 min to ensure that lipids and lipid-PEG conjugates completely covered the surface of the precipitated polymeric particles. Note that the amount of organic solvent used in this sonication method (10 vol %) was significantly less than what has been used in the previous nanoprecipitation method (25 vol %) that involves sample heating, vortexing, and solvent evaporation.25 This reduction of organic solvent allows one to forego the solvent evaporation step and skip straight to purification using centrifuge filters with a molecular weight cutoff of 10 kDa. When the weight ratios of lecithin/PLGA and DSPE-PEG/PLGA were 12.5 μg/ 1 mg and 200 μg/1 mg, respectively, this sonication method resulted in monodisperse lipid-PLGA hybrid nanoparticles with an average hydrodynamic diameter of 65 ( 1 nm and polydispersity index (PDI) of 0.08 ( 0.01 in PBS buffer (pH=7.4, 1X), determined by dynamic light scattering (Figure 1B). The morphology and low PDI of the hybrid nanoparticles were further confirmed by scanning electron microscopy (SEM) imaging, as shown in Figure 1C. The surface zeta potential of these nanoparticles, when dissolving in pure deionized water, was -47.7 ( 0.5 mV determined by DLS. However, it became difficult and inaccurate to measure the surface charge of the particles when they were suspended in 1X PBS because of the interference from high ionic concentration. In this work, the quick formation of lipid-coated polymeric nanoparticles may be attributed to the high energy and uniform means of energy input provided by bath sonication. In contrast to the conventional heating-vortexingevaporation method that provides for turbulent mixing and relatively low energy, this sonication approach will allow the building blocks to quickly complete kinetic self-assembly process 16960 DOI: 10.1021/la103576a
Figure 2. (A) Effect of the lipid-PEG/PLGA polymer weight ratio on nanoparticle size and polydispersity index (PDI). (B) Effect of lecithin/lipid-PEG molar ratio on the nanoparticle size and PDI. At a fixed lipid-PEG/PLGA weight ratio, a portion of the lipid-PEG was replaced by an equivalent mole amount of lecithin to prepare the hybrid nanoparticles. The size and PDI of the particles were determined by DLS in PBS buffer.
and result in a particle population with low polydispersity. This explanation is further supported by the fact that similar size nanoparticles can be achieved after 2-3 min of sonication, but PDI of the particles is not as small as those after 5 min sonication. In addition, sonicating for longer time will not affect particle size and PDI if all other environmental variables stay the same. Next, we examined the effects of relative concentrations of PLGA polymer, lipid, and lipid-PEG on the nanoparticle size and PDI. By using 2.5 mg/mL PLGA in acetonitrile solution and fixing the final aqueous to organic solution volume ratio of 10:1, we first explored the effect of lipid-PEG/PLGA weight ratio on particle formation and stability (Figure 2A). The mass ratio of lipid-PEG to PLGA polymer was varied from 0.10 to 0.35 while no lecithin was used to prepare the hybrid nanoparticles. The resulting nanoparticles were washed in PBS buffer, vortexed at 3000 rpm for 30 s, left at room temperature for 24 h, and then vortexed at 3000 rpm again prior to measuring their size and PDI. These rigorous operations were to test the stability of these nanoparticles as most of the unstable particles cannot tolerate PBS buffer or intense mechanical vortexing, which will induce particle aggregation. We found that more lipid-PEG in the formulation led to smaller particles and lower PDI. Especially, when the lipid-PEG/PLGA weight ratio was 0.30 (e.g., 300 μg of lipidPEG/1 mg of PLGA) or higher, particle size remained around 60 nm with a low PDI of about 0.09-0.10. This suggests that at such weight ratio the amount of lipid-PEG was sufficient enough to stabilize the PLGA particles. Langmuir 2010, 26(22), 16958–16962
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Figure 3. Long-term stability of the synthesized lipid-PLGA hybrid nanoparticles in terms of particle size and PDI in PBS buffer, which were monitored for a period of 5 days at room temperature.
We then fixed the lipid-PEG/PLGA weight ratio at 0.30 and replaced a fraction of lipid-PEG with the mole equivalent of lecithin. Particle size and PDI at various formulations were determined following the same protocol as described above. As shown in Figure 2B, the particles had an average size of about 60 nm and a PDI less than 0.10 when up to 50 mol % of lipidPEG was replaced by an equivalent molar amount of lecithin. However, large particles and particle aggregates form when more than 70 mol % of lipid-PEG was replaced by lecithin and the PDI of the particles increased dramatically as well. For example, at 80% replacement, the particle size increased to above 100 nm. At 100% replacement, the particles were unstable and visibly aggregated immediately in PBS. These studies suggest that the weight ratio of lipid-PEG/PLGA and the molar ratio of lecithin/ lipid-PEG are two effective factors to control the physicochemical properties of the hybrid nanoparticles. In the subsequent studies, we chose to use the formulation that has a lipid-PEG/ PLGA weight ratio of 0.30 with 33 mol % of the lipid-PEG replaced with its mole equivalent in lecithin. To evaluate the shelf stability of these lipid-polymer hybrid nanoparticles, we conducted particle stability tests in 1 X PBS buffer. The samples were kept at room temperature, and particle characteristics (size and PDI) were measured using DLS over a period of 5 days to monitor if aggregation would occur. As shown in Figure 3, no significant change in size or PDI was noted during the observation period, suggesting that this hybrid particle formulation could be stored for long periods of time with little or no aggregation. The particles remained at around 65 nm with a PDI of 0.08 over the course of the 5 days. Finally, we tested the serum stability of these hybrid nanoparticles, which would provide valuable information to evaluate the in vivo behavior of the particles. To monitor the particle size change at high concentration of serum (e.g., 50% serum or higher), we adapted a method that was previously used to test the serum stability of PEGylated polystyrene nanobeads by measuring changes in absorbance at a wavelength of 560 nm.31 Note that DLS is incapable of accurately measuring the size of sub-100 nm particles in dense serum solutions because of strong interference from the proteins and protein clusters in serum. PLGA nanoparticles at various sizes in deionized water were first used to confirm the particle size dependence of the absorbance intensity. Keeping the PLGA polymer concentration the same (1 mg/mL), four different-sized bare PLGA nanoparticles (diameter: 50, 69, 78, and 102 nm) were synthesized using a nanoprecipitation method with acetonitrile as the organic phase and pure water as the aqueous phase.17 Particle size was controlled by varying the initial PLGA concentration prior to nanoprecipitation. The Langmuir 2010, 26(22), 16958–16962
Figure 4. Serum stability of the synthesized lipid-PLGA hybrid nanoparticles. (A) Absorbance spectra of different sized bare PLGA nanoparticles (NPs) in water. The solution background (without nanoparticles) of each sample was subtracted to give the absolute particle absorbance. A sampling wavelength of 560 nm was confirmed to show a linear correlation between absorbance intensity and particle size at a constant total polymer weight. (B) Absorbance intensity at 560 nm of hybrid nanoparticles in 50% fetal bovine serum (FBS) over a period of 24 h. PLGA-PEG diblock copolymer nanoparticles in 50% FBS, bare PLGA nanoparticles in 50% FBS, and 50% FBS alone (without nanoparticles) were measured in parallel as controls. The background absorbance of water was measured and subtracted from each set of data.
absorbance spectra of the resulting samples were then measured, and the background absorbance of each solution (without nanoparticles) was subtracted to obtain absolute absorbance intensity of particles against particle size (Figure 4A). From the data obtained, we saw a clear correlation between the size of the particle and the strength of the absorbance signal. The graph confirmed that 560 nm was a suitable wavelength to use for monitoring particle size, although there were a wide range of wavelengths that we could have also used. For the actual serum stability tests, we examined four different samples: lipid-polymer hybrid nanoparticles in 50% FBS, PLGA-PEG diblock copolymer nanoparticles in 50% FBS as a negative control, bare PLGA nanoparticles in 50% FBS as a positive control, and 50% FBS alone (without nanoparticles) as a negative control. All samples were incubated at 37 °C with periodic gentle shaking. Absorbance values at the wavelength of 560 nm for each sample were taken over a period of 24 h, and the results were plotted against time with the water backgrounds subtracted (Figure 4B). We observed that lipid-PLGA hybrid nanoparticles had a similar absorbance value as that of PLGAPEG diblock nanoparticles, which was only slightly higher than that of 50% serum alone. These data suggest that there was no significant initial aggregation and that there was also no aggregation over time. In contrast, the bare PLGA nanoparticles visibly aggregated right away in 50% FBS, and the absorbance was DOI: 10.1021/la103576a
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already more than double the value of the serum baseline even before the first data point was taken.
Conclusions In conclusion, we have developed a new and fast sonication method to synthesize lipid-polymer hybrid nanoparticles that have sub-100 nm particle size and less than 0.10 polydispersity index. This method could be generalized to synthesize highquality hybrid nanoparticles within a wide range of relative concentrations of polymer, lipid, and lipid-PEG conjugate. The resulting nanoparticles were proven to have the potential to be stored long-term in PBS buffer solution. In addition, these
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particles did not aggregate when suspended in fetal bovine serum, indicating great promise to be used in vivo. As this new method does not involve the use of sample heating, vortexing, or solvent evaporation, it reduces the time needed to prepare the same amount of hybrid nanoparticles by a factor of about 20 without compromising the quality of particles. This approach makes it possible to scale up the production of lipid-polymer hybrid nanoparticles for possible preclinical and clinical tests in the future. Acknowledgment. This work is supported by the National Science Foundation Grant CMMI-1031239.
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