Formation of Monodisperse Hierarchical Lipid Particles Utilizing

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Formation of monodisperse hierarchical lipid particles utilizing microfluidic droplets in a non-equilibrium state Masahiro Mizuno, Taro Toyota, Miki Konishi, Yoshiyuki Kageyama, Masumi Yamada, and Minoru Seki Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.5b00043 • Publication Date (Web): 11 Feb 2015 Downloaded from http://pubs.acs.org on February 18, 2015

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Formation of monodisperse hierarchical lipid particles utilizing microfluidic droplets in a nonequilibrium state

Masahiro Mizuno,† Taro Toyota‡, Miki Konishi,† Yoshiyuki Kageyama,§ Masumi Yamada,*,† and Minoru Seki†

†

Department of Applied Chemistry and Biotechnology, Graduate School of Engineering, Chiba

University, 1-33 Yayoi-cho, Inage-ku, Chiba 263-8522, Japan. ‡

Department of Basic Science, Graduate School of Arts and Sciences, The University of Tokyo,

3-8-1 Komaba, Meguro, Tokyo 153-8902, Japan. §

Department of Chemistry, Faculty of Science, Hokkaido University, Kita 10, Nishi 8, Kita-ku,

Sapporo 060-0810, Japan.

*E-mail: [email protected]; Tel/fax: +81-43-290-3398.

ACS Paragon Plus Environment

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ABSTRACT

A new microfluidic process was used to generate unique micrometer-sized hierarchical lipid particles having spherical lipid-core and multilamellar-shell structures. The process includes three steps: (1) formation of monodisperse droplets in a non-equilibrium state at a microchannel confluence, using a phospholipid-containing water-soluble organic solvent as the dispersed phase and water as the continuous phase; (2) dissolution of the organic solvent of the droplet into the continuous phase and concentration of the lipid molecules; and (3) reconstitution of multilamellar lipid membranes and simultaneous formation of a lipid core. We demonstrated control of the lipid particle size by the process conditions and characterized the obtained particles by transmission electron microscopy and microbeam small-angle X-ray scattering analysis. In addition, we prepared various types of core-shell and core-core-shell particles incorporating hydrophobic/hydrophilic compounds, showing the applicability of the presented process to the production of drug-encapsulating lipid particles.


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INTRODUCTION The self-assembly of lipid molecules has drawn great attention not only as a tool for reconstituting the functions of lipid-associated biomolecules but also as a source of biocompatible vehicles for drug delivery systems. For instance, lipid aggregates incorporating hydrophobic compounds are thought to be a useful model of lipoprotein particles found in blood1,2. Unilamellar liposomes composed of closed bilayer membranes of lipids3,4, black lipid membranes5, and lipid monolayers formed at air/water or oil/water interfaces6,7 have been used as cellular membrane models. Multilamellar tubular liposomes structurally resemble the myelin sheath formed around the axon of a neuron8. Spherical nano/micrometer-sized lipid particles, or so-called solid lipid microspheres, as well as multilamellar liposomes, are promising candidates for drug delivery carriers1,9,10. These lipid structures can incorporate a variety of molecules; unilamellar liposomes are suitable for encapsulating hydrophilic compounds inside their aqueous cores11, whereas multilamellar liposomes can carry both hydrophobic and hydrophilic compounds within their intra-/inter-membrane structures, respectively12. Among the various physicochemical characteristics of lipid particles and liposomes, uniformity in size, as well as their internal structure, is one of the most important factors dominating the elution kinetics of encapsulated molecules, accumulation in the living body, and drug-loading capacity. To date, many approaches have been reported for the preparation of monodisperse unilamellar liposomes13-15.

Microfluidic technologies capable of efficiently

preparing monodisperse droplets16 have been applied in the production of unilamellar liposomes17,18. Uniformly sized liposomes with single lamella have been produced by generating aqueous droplets in an oil phase through the water/oil interface with the absorbed lipid


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molecules19, by swelling size-controlled lipid films20, or by using ice particles as an inner template21. In contrast, the direct production of monodisperse multilamellar liposomes is very difficult, and has not been fully developed. This may be attributed to the complexity of the process of forming such lipid particles or liposomes with multilamellar structures, which requires self-reconstitution of the lipid and water molecules into closely stacked bilayer membranes. One of the most frequently used methods to produce multilamellar liposomes is the swelling of dried lipid films with water, which results in the formation of liposomes with large size distributions14. In order to obtain a monodisperse population of multilamellar liposomes, specific techniques must be employed to select liposomes with desired sizes from a heterogeneous mixture, including filtration22, size exclusion chromatography23,24, and membrane dialysis methods25. Developments of several types of microfluidic technologies for preparing multilamellar lipid tubes have also been reported according to the hydration of accumulated lipid molecules within a confined microchannel26-28. However, these technologies require multistep, time-consuming operations; for example, the film-swelling method includes the deposition of phospholipid molecules, their swelling in an aqueous phase, collection, and size-based sorting. In the previous microfluidics-based techniques, the production efficiency and throughput for monodisperse lipid particles, liposomes, or tubes were limited.

A facile process for producing monodisperse

multilamellar lipid particles is of great importance. We recently reported strategies for producing monodisperse microparticles by using microfluidic droplets in a non-equilibrium state (i.e., non-equilibrium droplets)29,30. Oil-in-water (O/W) or water-in-oil (W/O) droplets were generated by using water and a polar organic solvent (e.g., ethyl acetate) as either the continuous or dispersed phase. The polymer molecules in the


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non-equilibrium droplets were gradually concentrated as the droplets were dissolved into the continuous phase, resulting in the formation of polymeric or hydrogel particles with sizes significantly smaller than those of the initially formed droplets. Moreover, we were able to control the particle morphologies by changing the degree of droplet dissolution and by controlling the segregation dynamics of the solid/liquid phases within the droplet29,30. This process, often referred to as “solvent diffusion”, has been utilized to produce various types of particles31; however, to the best of our knowledge, a process for producing lipid particles or liposomes using such non-equilibrium droplets has not been reported. In the present study, we propose a simple process for producing monodisperse hierarchical lipid particles from non-equilibrium droplets. The process is shown in Fig. 1. Droplets of a polar solvent (mainly ethyl acetate; EA) containing phospholipids are formed in the continuous water phase at a microchannel confluence, and gradually shrink while flowing through the microchannel because of solvent dissolution into the continuous phase. We expected that multilamellar liposomes would be generated by further dilution and complete removal of the solvent; however, we obtained hierarchical lipid particles having unique core-shell structures. We examined the sizes and morphologies of the core-shell lipid particles and evaluated their applicability as carriers for both hydrophobic and hydrophilic compounds.


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Figure 1. Schematic illustration showing the process of lipid microparticle preparation using a microfluidic device. EA: ethyl acetate. PBS: phosphate buffered saline.

MATERIALS AND METHODS Materials Lecithin (from egg), 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), ethyl acetate (EA, >99.9% purity), propyl acetate (>99% purity), diethyl ether (>99.5% purity), sodium fluorescein, rhodamine B, Nile red, sodium dodecyl sulphate (SDS), and olive oil were obtained from Wako Pure Chemical Industries Ltd., Osaka, Japan. Pyrene was obtained from Kanto Chemical Corp., Tokyo, Japan. Silicone oil (AR 200) and triton X-100 were purchased from Sigma-Aldrich Corp., MO, USA.

(Heptadecafluoro-1,1,2,2-tetradecyl)-trimethoxysilane and

phosphate buffered saline (PBS, Dulbecco′s formula) tablets were obtained from Gelest Inc., PA, USA and Takara Bio Inc., Shiga, Japan, respectively.

Preparation of core-shell lipid particles


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Polydimethylsiloxane (PDMS)-glass hybrid microdevices were fabricated using standard soft lithography and replica molding techniques as described elsewhere32. The microchannel design is shown in Fig. 2 (a). We employed a flow-focusing microchannel having an orifice structure to generate monodisperse O/W droplets. The width of the orifice was 50 µm and that of the main microchannel was 200 µm. The depth of the microchannel was approximately 100 µm. EA was used as the water-soluble organic solvent, unless otherwise noted. There were six inlets in this microdevice; one for the phospholipid solution in EA (Inlet 1), two for pure EA (Inlets 2 and 2′), and three for the continuous water phase (Inlets 3, 3′, and 4). The microchannel surface was modified by silanization: the PDMS and glass plates were bonded via O2 plasma treatment and subsequently silanized by introducing a small aliquot of 1% (heptadecafluoro1,1,2,2-tetrahydrodecyl)trimethoxysilane in methanol and incubating at room temperature for 5 min, just before conducting the microfluidic experiments. We used DOPC and lecithin as the phospholipid molecules dissolved in EA.

Lipid solutions were prepared by dissolving

phospholipid molecules in EA at concentrations varying between 1 and 5% (w/v). Distilled water or PBS (2.7 mM KCl, 1.5 mM KH2PO4, 136.9 mM NaCl, and 8.9 mM Na2HPO4•7H2O) was used as the continuous phase. The phospholipid EA solution, pure EA, and the continuous phase (distilled water or PBS) were continuously introduced into the microchannel by syringe pumps (KDS 200, KD Scientific, MA, USA). Inlet 4 was set to introduce additional continuous phase 500 µm downstream from the first confluence (orifice). The flow rates from Inlets 1, 2, 2′, 3, 3′, and 4 were 2, 2, 2, 20, 20, and 120 µL/min, respectively. The formed lipid particles were continuously collected via Teflon tube into a glass vial containing the continuous phase. The morphologies of the lipid particles were observed with an optical/fluorescence microscope (IX71, Olympus, Japan) with a CCD camera (DP72, Olympus), and sets of fluorescence excitation and


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detection filters (λex: 330–385 nm and λem: > 420 nm for blue dye; λex: 460–495 nm and λem: 510– 550 nm for green dye; and λex: 530–550 nm and λem: > 575 nm for red dye). The sizes of the cores and shells of the lipid particles and their distributions were obtained by measuring and averaging the longest and shortest axes of ~50 particles for each condition. For further analysis by transmission scanning electron microscopy (TEM, JEM-1400Plus, JEOL Ltd., Japan), the remaining EA was completely removed by exchanging the medium with the continuous phase several times. Then, the collected lipid particles were treated with 2% phosphotungstic acid solution (pH7.0) for 20 s (negative staining), and the microstructures were observed.

Microbeam SAXS measurements To elucidate the structure of the core-shell lipid particle, microbeam small-angle X-ray scattering (SAXS) measurements were performed at the BL-4A beamline in the Photon Factory (KEK, Ibaraki, Japan)33. After condensation of the lipid particles by centrifugation (2000 × g, 10 min, r.t.), the lipid particle dispersion was encapsulated in a chamber (5 mm × 5mm × 0.08 mm) made of two glass plates (thickness = 0.12–0.15 mm) and double-sided frame-type tape. The microbeam X-ray irradiation (10 keV), which was monochromated and focused to ~5 µm × 5 µm by Kirkpatrick-Baez mirrors, was scattered by the core or rim of the lipid particle and detected by a 1024 × 1024-pixel X-ray Image Intensifier CCD camera (C4880-50-26A, Hamamatsu Photonics, Japan). The scattering patterns were analyzed by Image J (NIH, USA) using a reference scattering pattern from a silver behenate sample obtained by the same X-ray irradiation protocol34. As a reference experiment, we used the microbeam SAXS patterns from micrometersized multilamellar liposomes which were prepared by agitating POPC powder in water (10 mM). To evaluate the Bragg peak positions, we used a Gaussian regression curve.


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RESULTS AND DISCUSSION Formation of non-equilibrium droplets and core-shell lipid particles We first examined whether the non-equilibrium EA droplets containing phospholipid would be formed in the microchannel. In general, hydrophilic surfaces should be employed to generate O/W droplets in microchannels. We initially tried to form droplets of phospholipid EA solution in O2-plasma-treated hydrophilic PDMS-glass microchannels. However, droplets were not generated; instead, the phospholipid EA solution flowed on the microchannel surface. To prevent wetting of the microchannel by the phospholipid EA solution and to stably generate the droplets, we modified the PDMS surface with the fluorosilane. In addition, we introduced pure EA between the phospholipid EA solution and the continuous water phase so that direct contact between the two phases was avoided; by this operation, the precipitation of lipid molecules near the confluence was prevented and the droplet formation became stable. Moreover, continuous phase was added into the microchannels from Inlet 4 to prevent the formed droplets from coalescing. Fig. 2 (b-d) shows the formation of EA droplets containing DOPC, using distilled water as the continuous phase and 1% (w/v) DOPC in EA. Monodisperse droplets of the phospholipid EA solution were generated at the first confluence with an average diameter of 89 µm and coefficient of variation (CV) value of 5.2%. The generated droplets gradually shrank while flowing through the microchannel, because of the dissolution of EA into the continuous aqueous phase. Near the exit of the microchannel, the droplet diameter decreased to 77 µm on average (CV, 2.8%) as shown in Fig. 2 d, where the retention time was ~0.4 sec. Droplet shrinkage was completed during flowing through the outlet Teflon tube (with the retention time of 10-20 sec).


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Fig. 3 (a, b) shows the formed DOPC lipid particles after collection from the microchannel into a glass vial. The lipid particles are highly monodisperse, and exhibit unique morphologies: a spherical core (average diameter, 12 µm; CV, 7.6%) and an outer shell around the lipid core (average diameter, 56 µm; CV, 18%). The shell of the particles was not uniform, incorporating small vesicle like-structures. To the best of our knowledge, the formation of lipid particles exhibiting such a core-shell morphology has not been reported. It should be noted that O/W droplets were not generated but multiphase parallel flows were formed when propyl acetate and diethyl ether were used as the polar organic solvent instead of EA. Additionally, it was also possible to produce similar core-shell lipid particles in a bulk-scale experiment, in which the phospholipid-EA solution was dropped into distilled water in a flask with vigorous stirring. However, the size and morphology of the obtained particles were not uniform and not reproducible, because the droplet formation and dissolution were not controllable.


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Figure 2. (a) Design of the microfluidic device with a main channel length of 40 mm. (b–d) Phase-contrast microscopy images of EA droplets containing DOPC in the microfluidic device. EA droplets with DOPC (b) generated at the first confluence point (orifice), (c) flowing through the second confluence point, and (d) flowing at 30 mm from the second confluence point.

Next, the continuous phase was changed from distilled water to PBS, to examine the effect of ionic strength on the morphology of the lipid particles. Under this condition, the shell size of the obtained lipid particles (22 µm; CV, 10%) was much smaller than that of the lipid particles prepared in water as the continuous phase, whereas the core size was nearly unchanged (11 µm; CV, 9.2%; Fig. 3 (c)). When we used 5 times denser PBS (5 × PBS) as the continuous


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phase, the particle size was almost unchanged but the relative size of the core was increased (shell diameter, 21 µm; CV, 10%; core diameter, 17 µm; CV, 14%; Fig. 3 (d)), indicating that the swelling of the shell was further suppressed. These results indicate that the swelling of the outer shell is suppressed when a continuous phase with a higher ionic strength is employed. Thus, the possibility of controlling the morphology of the lipid particles by using different continuous phases was shown. In addition, we controlled the size of the lipid particles by changing the initial phospholipid concentration in EA. Using DOPC concentrations of 1 or 5% and PBS as the continuous phase, the size distributions of the core-shell lipid particles were measured (Fig. 3 (e)). At the higher DOPC concentration, the core and shell diameters were 11 µm (CV, 8.8%) and 29 µm (CV, 7.7%), respectively. We confirmed that the shell size of the lipid particles increases when the DOPC concentration is higher, whereas the core sizes are nearly constant. These results show the possibility of controlling the particle size by changing the operating conditions (including the lipid concentration in the EA droplets, the size of the orifice at the device, etc.).


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Figure 3. (a-d) Phase-contrast micrographs of lipid particles prepared with (a, b) distilled water (under different magnifications), (c) 1 × PBS, and (d) 5 × PBS as the continuous phase. DOPC concentration in EA was 1%. (e) Core and shell size distributions of the obtained lipid particles at a DOPC concentration of 1% or 5%. 1 × PBS was used as the continuous phase.

Characterization of the lipid particles To characterize the obtained lipid particles, they were first introduced into aqueous surfactant solutions. Fig. 4 (a, b) shows the particles after introduction into an aqueous solution of 10 mM SDS. The shells of the particles dissolved, whereas significant changes were not observed for the cores in terms of shape and size. Similar results were obtained when 10% triton X-100 in distilled water was used as the surfactant solution. The cores of the particles were also separable from the shells by ultrasonication for 30 min (data not shown). These results indicate


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that the cores are most likely composed of condensed lipid molecules, as with solid lipid particles, whereas the shells were soft and fragile. Moreover, to examine the difference between the cores and shells, we dried the lipid particles on glass slides and swelled them in distilled water (Fig. 4 (c, d)). The shell structure was transformed into multiple spherical multilamellar liposomes with diameters of several micrometers, whereas the shape of the core was not significantly changed.

Figure 4. (a, b) Phase-contrast micrographs of the cores of the lipid particles isolated by treatment with 10% SDS solution. (c, d) Phase-contrast micrographs of (c) a dried core-shell lipid particle on a glass slide, and (d) re-swelled particle with distilled water.

TEM observations Next, we observed the microstructures within the core and shell of the lipid particles, which were prepared using distilled water as the continuous phase, by TEM with negative staining. As shown in Fig. 5 (a, b), multiple lipid bilayers can be clearly observed in the entire shell region of the lipid particles. The thickness of the bilayer is ~4.5 nm, which is very close to that of multilamellar DOPC liposomes prepared by the conventional film-swelling method3.


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This result indicates that the shells of the lipid particles are structurally identical to the multilamellar liposomes. This result was consistent with the aforementioned observation that the particle shell was easily dissolved after treating the hierarchical particles with aqueous surfactant solutions.

In contrast, we could not discern any microtextures within the core by TEM

observation, likely because of the randomly packed lipid molecules.

Microbeam SAXS analysis In the profiles of the SAXS scattering patterns from the rim of the lipid particle and the center of the POPC multilamellar liposomes, we observed the first Bragg peak in range of qm = 0.0984–0.1004 Å-1. Several higher-order peaks appeared at wave numbers equal to integral multiples of qm (Fig. 5 (c)). The SAXS data can be interpreted by the formation of a highly ordered lamellar structure having an interlayer distance, d, of 6.4 nm, where d = 2π/qm, at the rim of the lipid particle. It was also deduced that the POPC multilamellar liposomes with diameters of several tens of micrometers have an interlayer distance of 6.3 nm at the center. These results indicate that the rim of the lipid particle has a lamellar structure closely related to that of the center of a multilamellar liposome35; i.e., stacked bilayer membranes are formed as the shell structure at the rim of the lipid particle. We could not find any obvious peaks in the profile of the scattering pattern from the core of the lipid particle. These results correspond well with the TEM observations of the core-shell lipid particles. Judging from these results in concert with the TEM observations, we deduced that the obtained lipid particles in the current microfluidic process have a unique structure composed of a closely packed, non-ordered core and a multilamellar shell (Fig. 5 (d)). Additionally, we assumed that the particle core was mainly


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composed of the phospholipid molecules and did not contain EA, because the core morphology was maintained after the particle washing with distilled water and the drying/re-swelling process.

Figure 5. (a, b) Transmission scanning electron microscopy images showing the shell structures of the core-shell lipid particles. (b) is the enlarged image of the white rectangle in (a). (c) Microbeam SAXS analysis of the core-shell lipid particles. The red, dotted green, and dotted blue lines indicate the scattering patterns by the rim of the particles, center of the particles, and multilamellar POPC liposomes as control, respectively. (d) Schematic illustration of the coreshell hierarchical lipid particles.


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Generation behavior of core-shell lipid particles To further elucidate the formation mechanism of the core-shell lipid particles, we observed their generation behaviors. A longer microchannel (500 mm) was employed to extend the retention time to ~4 s (Fig. 6 (a)). Distilled water was used as the continuous phase, and the DOPC concentration in EA was 1%. The generation behaviors of the particles are shown in Fig. 6 (b)–(c). Approximately 1 s after droplet generation, the formation of lamellar shell structures was initiated around an inner EA droplet (Fig. 6 (c)). We assumed that thick membranes were formed at the interface between the continuous phase and the shrinking droplet, which were generated by the EA and DOPC diffused from the shrinking inner droplet and water molecules supplied from the continuous phase. The shell structures continuously grew around the shrinking droplets and gradually became thicker (Fig. 6 (d)–(f)). This indicates that the DOPC and water molecules are reconstituted into multilamellar structures as EA is gradually dissolved into the continuous phase.

Droplet shrinkage and formation of the multilamellar shell were nearly

complet 3.5 s after droplet generation (Fig. 6 (g)), forming solid core structures, which were probably generated by the precipitation and segregation of DOPC molecules.

From these

observations, we confirmed that complex condensation processes of the DOPC molecules occur in the droplet and at the interface between the continuous and dispersed phases, resulting in the formation of the core-shell hierarchical lipid particles.


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Figure 6. (a) Design of a microfluidic device with a longer (~500 mm) main channel. (b-g) Phase-contrast micrographs of droplets/particles at different sites along the microchannel, with corresponding retention times. Note that different droplets/particles are shown for these images.

Preparation of various types of lipid particles Because this process is operationally simple and reproducible, we anticipated the possibility of producing lipid particles with different compositions. We first examined the applicability of the presented process in the production of different lipid particles types by using lecithin as the lipid. Lecithin is a mixture of several types of phospholipid molecules including


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phosphatidylcholines (~ 70%) and phosphatidylethanolamines (~15%), in which the two acyl substituents at positions 1 and 2 are specified as palmitoyl (~35%), oleoyl (~30%) and stearoyl (~10%)36,37. The lipid particles obtained using distilled water as the continuous phase and 1% lecithin in EA as the dispersed phase are shown in Figure 7. Lipid particles with core-shell morphologies similar to those of the DOPC particles were obtained even though lecithin is a complex lipid.

Figure 7. Phase-contrast micrographs of monodisperse core-shell lipid particles produced by lecithin from egg yolk as the lipid molecule. Image (b) is a magnified view of a particle in (a).

It is likely that one could encapsulate a variety of molecules within them, because the presented lipid particles are composed of two distinct parts, a multilamellar shell and a condensed lipid core. To validate this concept, we next prepared lipid particles incorporating hydrophobic and hydrophilic compounds.

First, hydrophobic dyes, including Nile red,

rhodamine B, and pyrene were encapsulated. These molecules were individually dissolved in the dispersed phase of EA containing 1% DOPC, at molar ratios of 1, 5, and 1% with respect to DOPC, respectively. The lipid particles obtained using distilled water as the continuous phase are shown in Fig. 8 (a-f). The hydrophobic molecules were encapsulated within the shells of the particles and stably retained even after rinsing with the continuous phase.


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The multilamellar shells of the lipid particles were composed of lipid bilayers, which were formed by the supply of water molecules from the continuous phase. We therefore tested whether hydrophilic compounds, initially dissolved in the continuous phase, would be incorporated into the lipid particles. Sodium fluorescein was added into the continuous phase (distilled water) at a concentration of 0.4 µg/mL. Fig. 8 (g, h) shows the obtained lipid (DOPC) particles. Green fluorescence was observed from the shell and the boundary region between the shell and core of the particles. This result indicates that the hydrophilic molecules were also incorporated into the shells of the lipid particles together with water at the time of droplet dissolution and shell formation. No fluorescence was observed from the core, which was confirmed by dissolving the shell in 10 mM aqueous SDS solution. These results demonstrate the ability of the lipid particles to encapsulate both hydrophilic and hydrophobic molecules, indicating their versatility compared with unilamellar vesicles.

In addition, multilamellar

liposomes have been used as carriers for gene delivery to living cells38 and as synthetic vaccines encapsulating protein antigens39, and the presented particles might also be applicable as carriers for such biomolecules. It should be noted that the encapsulated hydrophobic/hydrophilic dye molecules were stably retained in the lipid particles and did not leak from the particles even after incubating 1 week in distilled water at room temperature. We expect that the encapsulated molecule would likely be released by the enzymatic degradation of the phospholipid molecules, when the presented lipid particles are injected in the living body. Finally, we prepared lipid particles containing hydrophobic liquid molecules that are soluble in EA, insoluble in water, and unable to solubilize phospholipid molecules. Such liquid samples were expected to be concentrated in the droplets as they dissolved and finally form independent droplets. To examine the effects of such liquid molecules on the morphology of the


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lipid particles formed, we employed silicone oil and olive oil, which were dissolved in EA at 0.5 and 1.0%, respectively, together with 1% DOPC. Distilled water was used as the continuous phase, and lipid particles were prepared.

To visualize the different parts of the particles,

rhodamine B was added to the phospholipid EA solution (0.3 mol% relative to DOPC). The obtained particles are shown in Fig. 8 (i-l). When silicone oil was added, ~40% of the obtained particles showed a unique core-core-shell morphology (Fig. 8 (i, j)), whereas the rest (~60%) were similar to those shown in Fig. 8 (c, d). The inner core exhibited the strong red fluorescence of rhodamine B, indicating that this core was mainly composed of silicone oil. It is likely that this inner core was formed by the segregation of a condensed lipid particle and a droplet of silicone oil during the dissolution process, but some of the particles might have lost the inner core during dissolution. This kind of hierarchical, liquid droplet-encapsulated phospholipid particle has not been reported yet, but it may be potentially useful as a new type of vehicle for controlled drug delivery and elution. In addition, we found that lipid particles with different morphologies were generated using different types of oils; twin cores composed of an oil phase and a phospholipid particle were observed in the shell, when olive oil was used (Fig. 8 (k, l)). This difference might be caused by the physicochemical characteristics of the oil, including the interfacial tension and the contact angle with the continuous or dispersed phase.


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Figure 8. Various types of core-shell lipid particles prepared by using DOPC as the lipid molecule. Phase-contrast and fluorescence micrographs of (a, b) a particle prepared by adding Nile red to the dispersed phase; (c, d) a particle prepared by adding rhodamine B to the dispersed phase; (e, f) a particle prepared by adding pyrene to the dispersed phase; and (g, h) a particle prepared by adding sodium fluorescein to the continuous phase. Phase-contrast and fluorescence micrographs of (i, j) a particle prepared by adding silicone oil and rhodamine B to the dispersed phase; and (k, l) a particle prepared by adding olive oil and rhodamine B to the dispersed phase.


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CONCLUSIONS A simple microfluidic system was developed to prepare unique monodisperse hierarchical lipid particles having lipid-core and multilamellar-shell structures, which are advantageously used for effective hydrophobic/hydrophilic molecule encapsulation.

The

presented lipid particles may be useful as functional lipid-based materials or vehicles for pharmaceutical, food, and cosmetic applications, and would be available as high-performance carriers for controlled drug delivery because we can precisely control the particle size and easily conduct surface modification or functionalization by incorporating functional molecules such as antibodies or magnetic particles in the multilamellar shells. Further control of the particle size and morphologies would be possible by changing the initial droplet size and the compositions of the continuous and dispersed phases.

ACKNOWLEDGMENTS This study was supported in part by KAKENHI (23106007 and 25103009) from MEXT, Japan. The microbeam SAXS analysis has been performed under the approval of the Photon Factory Program Advisory Committee (Proposal no. 2012G747).


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TABLE OF CONTENTS GRAPHIC


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Formation of Monodisperse Hierarchical Lipid Particles Utilizing

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