Osmotic-Stress-Mediated Control of Membrane Permeability of

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Osmotic-Stress-Mediated Control of Membrane Permeability of Polymeric Microcapsules Sangmin Lee, Tae Yong Lee, Dong Jae Kim, Bomi Kim, and Shin-Hyun Kim Chem. Mater., Just Accepted Manuscript • DOI: 10.1021/acs.chemmater.8b03230 • Publication Date (Web): 27 Sep 2018 Downloaded from http://pubs.acs.org on September 28, 2018

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Chemistry of Materials

Osmotic-Stress-Mediated Control of Membrane Permeability of Polymeric Microcapsules Sangmin Lee†, Tae Yong Lee†, Dong Jae Kim†, Bomi Kim†, and Shin-Hyun Kim*,† †

Department of Chemical and Biomolecular Engineering, Korea Advanced Institute of Science and Technology (KAIST), Daejeon, 34141, Republic of Korea ABSTRACT: Microcapsules with regular pore size can provide size-selective permeation, which is promising for immunoisolation of cells, protection of enzymes or catalysts, and development of capsule-type sensors. However, conventional approaches have limited biocompatibility or poor dispersion stability of encapsulants. Here, we suggest a simple yet pragmatic method to produce semipermeable microcapsules using osmotic stress. With a capillary microfluidic device, monodisperse microcapsules with ultra-thin polymer membranes are prepared by double-emulsion templating. The microcapsules are subjected to a hypotonic condition, by which water is pumped in, imposing a tensile stress on the membrane. The osmotic stress initiates cracks at weak spots. As cracks propagate, the pressure gradually reduces as ions diffuse through them, finally resulting in a finite width of cracks. The final width can be controlled from 5 to 10 nm using an initial osmotic pressure of 230 to 690 kPa, enabling fine adjustment of the cut-off threshold of permeation. This osmotic-pressure-mediated control is highly compatible with delicate biological molecules and colloidal dispersions as no etching chemicals are required to form pores. Taking advantage of this method, we demonstrate a capsuletype molecular sensor based on surface-enhanced Raman scattering that obviates pretreatment of samples because the membrane allows the entrance of small target molecules while blocking the large adhesive proteins.

INTRODUCTION Microcapsules are microcompartments encompassed by solid membranes, which are designed to provide desired functionalities.1-4 For example, dense membranes do not allow transmembrane transport, which is useful for long-term isolation of encapsulant materials such as chemical pigments and photonic structures.5-8 Alternatively, porous membranes provide controlled material exchange between the compartment and its surroundings.9-15 Membranes that have external stimulidependent permeability or degradation rates are promising for triggered release of chemicals or bio-actives on-demand.16-21 Such microcapsules are typically prepared by layerby-layer (LBL) deposition of polyelectrolyte on colloidal templates,22,23 interfacial polymerization,24-26 or particle adsorption at emulsion interfaces.27,28 However, these conventional methods are applicable only for limited sets of membrane materials and frequently suffer from low efficiency of encapsulation. As an alternative, techniques utilizing droplet microfluidics may enable the controlled production of multiple-emulsion drops, which serve as versatile templates to design microcapsules.29-31 In-situ loading of encapsulants during the droplet formation in the microfluidic device significantly improve the efficiency of encapsulation and the intrinsic core-shell geometry of the multiple-emulsion drops expands the sets of materials applicable to form membranes. Therefore, the microfluidics-based approach allows for a wide variety of membrane properties and functionality of capsules. Microcapsules with size-selective permeability have great potential in a wide range of applications, including immunoisolation of cells,15,32 protection of catalysts or enzymes,33-35 and as molecule-selective sensors.13,19,36,37 To pre-

pare the semipermeable microcapsules, double-emulsion templates have been microfluidically generated and employed in previous work to make membranes with a consistent size of pores. For example, nanoparticles are aggregated in the shell layer by solvent evaporation, which results in regular pores along the interstitial voids of the nanoparticle aggregates;2,12,38,39 however, the low mechanical stability of the membranes restricts their use. In other examples, polymerization-induced macro- or micro-phase separation have been used to make regular pores in stable membranes;9-11,40 however, selective removal of one phase requires a chemical treatment, which can reduce the dispersion stability of encapsulants or be lethal for biomolecules or cells. As an alternate approach, monomers are partially polymerized in the shell by controlling ultraviolet (UV) dose;41 however, it is difficult to uniformly expose the entire shell to the UV due to their spherical geometry. Others have used a mixture of monomer and volatile solvent as an oil shell,13 but it is difficult to control the cut-off threshold of permeation with this approach. Recent work as demonstrated formation of a hydrogel membrane by using water-in-water-in-oil (W/W/O) double-emulsion drops, which provides size-selective permeability without losing the dispersion stability of encapsulants;42 however, rapid polymerization of the monomer in water shell is required as the two aqueous phases are miscible. Therefore, it remains an important challenge to design semipermeable microcapsules with a high mechanical stability and controllable cut-off threshold of permeation through a benign approach compatible with various encapsulants. Here, we report a simple yet useful method to design semipermeable microcapsules with a controlled cut-off threshold. With a glass capillary microfluidic device, doubleemulsion drops are prepared to have an aqueous core and ul-

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tra-thin shell layer of an organic solution of biodegradable polymers. The microcapsules are prepared by consolidating the biodegradable polymer through solvent evaporation. When the microcapsules are stressed under hypotonic conditions, the membranes turn permeable to small molecules. The osmotic stress leads to the formation of cracks along the ultra-thin membranes, through which molecules are transported. The cut-off threshold of permeation, or width of cracks, can be controlled in the range of 5 – 10 nm by adjusting the osmoticpressure difference between the core and surrounding. As a physical stress is only applied along the membrane without chemical stress, high dispersion stability of encapsulants and low toxicity can be ensured. Using the microcapsules with size-selective permeability, we demonstrate a capsule-type molecular sensor by encapsulating aggregates of gold nanoparticles. Due to irreversible adsorption of protein on to the gold nanoparticles, which preclude access of small target molecules, it is hard to get adequate Raman signal without filtering process. However, the osmotic-pressure-driven membrane is designed to allow the infusion of small target molecules while excluding large proteins. Therefore, the Raman spectrum of small molecules is selectively amplified by the aggregates in the core through surface-enhanced Raman scattering (SERS) without interruption of proteins.

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thickness of the shell is approximately 0.9 µm which is estimated from the diameter of the ruptured drops, as shown in Figure S1. When the toluene solution passes through the tip, large oil drops are formed. Therefore, W/O/W doubleemulsion drops and O/W single-emulsion drops are cyclically produced. As most of the toluene solution forms oil drops and only a small part is incorporated into the shells of doubleemulsion drops, the shell thickness hardly changes with flow rates in the device with a discontinuous core-sheath flow.44 The emulsion drops are collected in an aqueous solution of 2% w/w PVA and 170 mM NaCl, which has the same osmolarity to the aqueous solution used for the innermost phase but has a lower density. The double-emulsion drops sink in the collection liquid, whereas single oil drops float, enabling the autonomous separation. The double-emulsion drops are incubated for 4 hours at room temperature in an open dish to evaporate the toluene, which fully consolidates PLGA in the shells.

RESULTS AND DISCUSSION Microfluidic production of microcapsules with ultra-thin polymer membrane. Microcapsules composed of an aqueous core and ultra-thin polymer membrane are prepared by using water-in-oil-in-water (W/O/W) double-emulsion droplets as a template.43,44 With a capillary microfluidic device, we produce double-emulsion drops with uniform size and composition, as shown in Figure 1a, b, and Movie S1. The capillary device is composed of two tapered cylindrical capillaries that are coaxially assembled in a square capillary to have a tip-to-tip configuration. One of the tapered capillaries has an orifice of 100 µm and hydrophobic walls and the other has an orifice of 180 µm and hydrophilic walls, where the tip-to-tip separation is 115 µm. Through the hydrophobic capillary, two immiscible phases— an aqueous solution of 5% w/w poly(vinyl alcohol) (PVA) and 150 mM sodium chloride (NaCl) and a toluene solution of 10% w/w poly(lactic-co-glycolic acid) (PLGA)—are simultaneously injected; the aqueous solution has osmolarity of 360 mOsm/L. Due to the hydrophobic nature of the capillary walls, the toluene solution flows along the wall, whereas the aqueous solution flows through the center of the capillary without contacting the wall. The aqueous solution forms plug-like drops rather than a jet as the biphasic flow is not strongly confined by the solid wall due to large radial dimension.44,45 This discontinuous core-sheath flow is emulsified into an aqueous solution injected through the gaps between the hydrophobic and square capillaries; the aqueous solution with the same composition to the one injected through the hydrophobic capillary is used as the continuous phase. The continuous phase exerts a strong drag force on the core-sheath stream flowing out from the tip of the hydrophobic capillary due to the flowfocusing effect, resulting in the breakup of the stream and formation of double-emulsion drops. The thinness and low flow rate of the oil sheath in the hydrophobic capillary make the oil shell of the double-emulsion drops extremely thin. The

Figure 1. (a) Schematic illustration of a capillary microfluidic device for preparation of double-emulsion drops with ultra-thin shell, where hydrophobic capillary is illustrated in red and the hydrophilic capillary in blue. (b) Still-shot optical microscopy (OM) image captured by a high-speed camera showing the formation of water-in-oil-in-water (W/O/W) double-emulsion drops. (c) Cartoons showing the preparation of ultra-thin shelled microcapsules by the consolidation of PLGA in toluene shell (first step) and the formation of cracks on the consolidated shell by osmotic stress at the hypotonic condition (second step). (d) The relative radius of microcapsules osmotically-stressed to original ones as a function of osmolarity difference between the core and surrounding, ∆C = Ccore – Csur. The relative radius remains unchanged for ∆C in the range between –40 and 300 mOsm/L. (e) OM image of monodisperse microcapsules stressed at ∆C = 100 mOsm/L.

The resulting microcapsules are highly monodisperse as shown in Figure S2a. The formation of the solid membrane


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is confirmed by squeezing the resulting microcapsules with a pair of glass plates; liquid shells rupture and form oil drops, whereas solid membranes deflate while maintaining their integrity, as shown in Figure S2b. The thickness of shell membrane decreases during the consolidation to approximately 145 nm, as shown in Figure S2c. As the PLGA polymers are densely packed during consolidation, the membranes do not allow diffusion of small dye molecules of sulforhodamine B (SRB, Mw 558.666 g/mol) as shown in Figure S2d. Influence of tonicity on microcapsules. The PLGA membranes are permeable for water, while impermeable for salt ions of sodium and chloride as well as PVA. Therefore, the osmotic pressure difference between the aqueous core and the external surroundings causes a flow of water through the membrane. When the osmotic pressure in the core is higher than that in the surrounding, water is pumped in, imposing a tensile stress on the spherical membrane; such conditions are referred to as hypotonic. The stress possibly produces cracks or pores on the membranes made of brittle PLGA, as illustrated in Figure 1c. It has been reported that microcapsules release encapsulants when they are subjected to hypotonic conditions.21,30,46,47 To study the permeability of osmotically-stressed membranes, the microcapsules are suspended in aqueous solutions of PVA and NaCl with different osmolarities of 60, 160, 260, 360 and 400 mOsm/L for 1 day, where a concentration of PVA is set to 5% w/w same to the core for all solutions and that of NaCl is varied. As the core has an osmolarity of 360 mOsm/L, the microcapsules suspended in the solutions with osmolarities of 260, 160, and 60 mOsm/L are in hypotonic condition with ∆C = 100, 200, and 300 mOsm/L and osmotic pressure differences (Π = ∆CRT) of 230, 460, and 690 kPa, where R [L kPa K−1 mol−1] is a gas constant and T [K] is temperature. The microcapsules do not show any noticeable physical change under observation with optical microscopy, as shown in Figure S3a-c. The microcapsules remain spherical and the radius is unvaried, as shown in Figure 1d, e. This is in stark contrast to elastomeric membranes which expand until the osmotic pressure difference is balanced by the elasticity of the membrane.48 When sulforhodamine B is additionally dissolved in the aqueous solutions, it infuses into the core, indicating that the membrane has cracks or pores, of which dimension is beyond the resolution of optical microscopy. In contrast, the microcapsules suspended in the aqueous solution with ∆C = 0 (isotonic condition) do not allow the infusion of dye, as shown in Figure S2d. The microcapsules suspended in the solution with ∆C = –40 mOsm/L (Π = –94 kPa, hypertonic condition) experience inward buckling of the membrane while maintaining the radius as water is pumped out, as shown in Figure S3d; although isotropic shrinkage of microcapsule is expected before buckling, the size reduction is insignificant for capsules with small membrane thickness relative to its radius.49 To verify the formation of cracks or pores at the nanoscale, the microcapsules with ∆C = 300 mOsm/L are completely dried and their membranes are inspected using scanning electron microscopy (SEM), as shown in Figure S4. The microcapsules are coated with 2-nm-thick Osmium tetroxide before inspection in order to resolve crack widths on the nanometer scale. As the membrane of microcapsules is composed of entangled PLGA, high mechanical stability is ob-

tained after drying process. At high magnification, crack-like features are observed as denoted with arrows. Although craters with a diameter of approximately 200 nm are also observed, it turns out that there is no transmembrane transport between the core and surrounding through these craters. In fact, the craters are observed in the microcapsules without osmotic stress. At hypotonic condition, the inward flux of water driven through the membrane by osmosis pressurizes the core, causing a tensile stress on the membrane. The natural outcome of tensile stress on a brittle polymer such as PLGA is crack formation rather than pore formation, as expected by Griffith theory of brittle fracture. Control over the cut-off threshold of permeation by osmotic pressure. As the ultra-thin membrane is made of brittle PLGA, the tensile stress is able to initiate cracks at weak spots, which propagate along the membrane over time. The tensile stress caused by osmotic pressure difference with ∆C = 25 mOsm/L is confirmed to be sufficient to create cracks in the membrane based on transmembrane transport of dye molecules, as shown in Figure S5. It is expected that the cracks do not open very widely because osmotic pressure difference decreases as sodium and chloride ions rapidly diffuse once the cracks form. Nevertheless, the width of the cracks depends on the initial osmotic pressure difference. To study the influence of the pressure difference on crack opening and transmembrane transport behavior, the osmotically stressed microcapsules are suspended in aqueous solutions of dye-tagged dextran molecules with different molecular weights; either fluorescein isothiocyanate (FITC)- or rhodamine B isothiocyanate (RITC)-tagged dextrans are used to visualize the diffusion of the molecules. The dextran molecules whose hydrodynamic diameter is smaller than the crack width diffuses through the membranes to the core, while those larger than the width are excluded from the microcapsules, as illustrated in Figure 2a. Most of microcapsules stressed by ∆C = 100 mOsm/L (Π = 230 kPa) allow infusion of dextran with Mw of 10,000 g/mol (hydrodynamic diameter, dH = 4.6 nm), while excluding that with Mw of 20,000 g/mol (dH = 6.6 nm), as shown in the left column of Figure 2b.50 From the hydrodynamic diameters, the cut-off threshold of permeation for the microcapsules is roughly approximated as 5.6 nm. Microcapsules stressed by ∆C = 200 mOsm/L (Π = 460 kPa) is permeable to the dextran with Mw of 20,000 g/mol and impermeable to that with Mw of 40,000 g/mol (dH = 9 nm) as shown in the middle column, indicating the cut-off threshold is roughly 7.8 nm. Microcapsules stressed by ∆C = 300 mOsm/L (Π = 690 kPa) is permeable to the dextran with Mw of 40,000 g/mol and impermeable to that with Mw of 70,000 g/mol (dH = 12 nm) as shown in the right column, indicating the cut-off threshold is about 10.5 nm. The fraction of microcapsules that allow the diffusion of dye molecules with various molecular weights and hydrodynamic diameters are summarized in Figure 2c; microcapsules with a fluorescent intensity larger than 30% of that of its surroundings is considered permeable. The microcapsules with ∆C = 0 do not allow the diffusion of all the dyes employed as no osmotic stress is applied; there is an infusion of SRB into less than 8% of microcapsules, which is attributed to intrinsic defects formed during the microcapsule preparation. All microcapsules stressed by ∆C = 100 mOsm/L allow the diffusion of SRB and the fraction decreases to 90% for dextran



with Mw of 4,000 g/mol (dH = 2.8 nm), 61% for Mw of 10,000 g/mol (dH = 4.6 nm), and 15% for Mw of 20,000 g/mol (dH = 6.6 nm). The fraction for the microcapsules stressed by ∆C = 200 mOsm/L is 100% for SRB, 75% for dextran with Mw of 10,000 g/mol (dH = 4.6 nm), 60% for Mw of 20,000 g/mol (dH = 6.6 nm), and 4% for Mw of 40,000 g/mol (dH = 9 nm). Microcapsules stressed by ∆C = 300 mOsm/L show higher fractions than those by ∆C = 200 mOsm/L for most infusing molecules. The fraction is 87% for dextran with Mw of 20,000 g/mol (dH = 6.6 nm), 76% for Mw of 40,000 g/mol (dH = 9 nm), and 0% for Mw of 70,000 g/mol (dH = 12 nm). We can further confirm that the craters with a diameter of 200 nm on the membrane do not allow transmembrane transport between the core and surroundings of the microcapsules as the cut-off threshold is much smaller than the diameter. The values of dH at the fraction of 50% are roughly 5, 7, and 10 nm for ∆C = 100, 200, and 300 mOsm/L, respectively.

Figure 2. (a) Schematic of a microcapsule with consistent crack size on the membrane which allows infusion of molecules smaller than crack width, while rejecting molecules larger than the width. (b) Confocal laser scanning microscopy (CLSM) images for three different microcapsules stressed by ∆C = 100 (left column), 200 (middle column), and 300 mOsm/L (right column). The microcapsules are suspended in an aqueous solution of FITC-tagged dextran with molecular weight denoted in each panel. The microcapsules are mostly permeable in top panels and mostly impermeable in bottom panels. (c) The fraction of molecule-permeable microcapsules as a function of hydrodynamic diameter (bottom xaxis) or molecular weight of dextran (top x-axis) for four different microcapsules including ones with no osmotic stress (∆C = 0). (d) CLSM images of microcapsules stressed by ∆C = 100 mOsm/L suspended in a binary aqueous solution of sulforhodamine B (SRB) and FITC-tagged dextran with molecular weight of 20,000 g/mol (top row) and microcapsules stressed by ∆C = 200 mOsm/L suspended in a binary aqueous solution of RITC-tagged dextran with molecular weight of 10,000 g/mol and FITC-tagged dextran with molecular weight of 40,000 g/mol (bottom row). Third panels in both rows are overlaid images.

To further prove the size-selective permeability, osmotically-stressed microcapsules are suspended in an aqueous solution containing two molecules of different sizes. When the microcapsules stressed by ∆C = 100 mOsm/L is suspended in

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the binary solution of SRB and FITC-tagged dextran with Mw of 20,000 g/mol, SRB diffuses into the microcapsules, whereas the dextran is rejected, as shown in the top row of Figure 2d. As we studied, the microcapsules stressed by ∆C = 100 mOsm/L have a cut-off threshold of 5 nm in diameter which is in between dH values of SRB and dextran with Mw of 20,000 g/mol. When the microcapsules stressed by ∆C = 200 mOsm/L is suspended in the binary solution of RITC-tagged dextran with Mw of 10,000 g/mol and FITC-tagged dextran with Mw of 40,000 g/mol, RITC-dextran diffuses into the microcapsules, whereas FITC-dextran is rejected, as shown in the bottom row of Figure 2d. We further confirm that the cracks on the membrane allow the release of encapsulated molecules to external surroundings through the following experiment. Here, FITCtagged dextran with Mw of 10,000 g/mol is encapsulated in the microcapsules, as this has been shown to permeate through the membranes into the microcapsules for all osmoticallystressed conditions. We can thus use this to also confirm release from the microcapsules to the external environment. The microcapsules are subject to hypotonic conditions with ∆C = 0, 34, 263, and 500 mOsm/L, respectively. The microcapsules with ∆C = 0 mOsm/L show negligible leakage of FITCdextran for at least two weeks, as shown in Figure S6a. By contrast, the microcapsules with ∆C = 34 mOsm/L release half of the encapsulants within 74 hours and those with ∆C = 263 mOsm/L do the same within 45 hours. When the osmolarity difference increases to ∆C = 500 mOsm/L, the rate of release does not significantly increase in a comparison with ∆C = 263 mOsm/L. This is because the pressure difference gradually decreases as sodium and chloride ions rapidly diffuse through the cracks. That is, the width of cracks is not linearly proportional to the initial pressure difference but gradually saturates. Permeability of osmotically-stressed microcapsules. The microcapsules stressed with ∆C = 100, 200, and 300 mOsm/L are all permeable to SRB. However, the rate of permeation is expected to be different as the size of cracks depends on the osmotic pressure difference. To characterize the rate, the microcapsules are suspended in an aqueous solution of SRB and the temporal change of fluorescent intensity of SRB in the core of microcapsules is recorded, as shown in Figures 3a and b. As the SRB diffuses through the cracks on the membrane into the core, the fluorescent intensity of microcapsules gradually increases until being comparable to that of the surrounding. It is clearly shown that the diffusion rate of SRB is higher in microcapsules stressed with ∆C = 300 mOsm/L than that in microcapsules stressed with ∆C = 100 mOsm/L. To quantitatively extract the timescale for diffusion, the increase of the intensity from the initial, I(t) – I0, is normalized by the difference between the maximum and initial values, I∞ – Io, of which temporal change is plotted for three different microcapsules, as shown in Figure 3c. In all three microcapsules, the normalized intensity rapidly increases in the early stage and then slows down, finally reaching a plateau. This temporal change follows the diffusion equation and a characteristic timescale, τ, is extracted from fits with the simplified solution of Fick’s law: ூሺ௧ሻିூబ ூಮ ିூబ

೟

= 1 − ݁ ିഓ .

(1)

The values of τ for the microcapsules stressed by ∆C = 100, 200, and 300 mOsm/L are 480, 339, and 170 minutes, respec-


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tively. From τ = r/3P and D ~ Pl, the values of D are approximated as 1.53, 2.17, and 4.32 × 10-16 m2/s respectively, where r is the radius of microcapsules (= 91.5 µm), P permeability, l thickness of membrane (= 145 nm) and D diffusivity. The free diffusion coefficient of SRB in 5% PVA solution is estimated as 8.7 × 10-11 m2/s from Stokes-Einstein equation. The diffusivity of SRB along the microcapsule membranes is five orders of magnitude smaller than the free diffusivity. This is because cracks take a very small areal fraction of entire surfaces of the membrane and the opening is only one order of magnitude larger than the hydrodynamic diameter of SRB.

Figure 3. (a, b) Time series of CLSM images showing an infusion of sulforhodamine B from the surroundings for two different microcapsules stressed by (a) ∆C = 100 and (b) 300 mOsm/L. (c) Temporal change of normalized fluorescence intensity for three different microcapsules stressed at ∆C = 100, 200, and 300 mOsm/L, where I0 and I∞ indicate initial and steady-state fluorescence intensities, respectively.

Microcapsules with size-selective permeation for molecular sensors. The microcapsules with size-selective permeability are promising as a microcontainer of molecular sensors. As the microcapsules retain sensing materials larger than the cut-off threshold in the core without a leakage, the sensing materials are not diluted by sample fluids. More importantly, the microcapsules reject molecules larger than the cut-off threshold while allowing infusion of small molecules. Therefore, adhesive large proteins which commonly interfere with sensing of the desired molecules by irreversible adsorption to the sensing materials can be excluded from the microcapsules and sensing materials remain intact. We encapsulate aggregates of gold nanoparticles in the microcapsules and produce cracks on the membrane by imposing ∆C = 100 mOsm/L at hypotonic condition to design molecular sensors based on surface-enhanced Raman scattering (SERS), as shown in Figure 4a; the aggregates of gold nanoparticles provide much higher enhancement of Raman signal than individual nanoparticles due to the formation of hot spots at the interstitial voids in the aggre-

gates.51,52 The aggregates of gold nanoparticles are prepared by chelating citrate groups of gold nanoparticles with copper ions according to a protocol previously reported, as shown in Figure S7a;53 it is known that the gold nanoparticles form twodimensional sheets. The aggregates are uniformly encapsulated in the microcapsules, as shown in Figure 4b.

Figure 4. (a) Schematic showing size-selective infusion of small molecules and enhancement of Raman scattering of the molecules by aggregates of gold nanoparticles in the microcapsules. (b) OM image of monodisperse microcapsules containing aggregates of gold nanoparticles. Inset is an OM image that brings the aggregates into focus. (c) Raman spectra of rhodamine 6G (R6G) measured from the central part of the aggregates in the microcapsules that are suspended in aqueous solution for the concentration range of 10-3 to 10-9 M. (d) Raman intensities at three different peak positions of 613, 773, and 1510 cm-1 as a function of the concentration. (e) CLSM images of microcapsules suspended in a binary aqueous solution of R6G and FITC-tagged BSA. (f) Raman spectra measured from the central part of aggregates in the microcapsules suspended in 0.1 mM R6G (blue line) and microcapsules in 0.1 mM R6G and 1 mM BSA (red line), which are comparable in the intensity. Raman spectrum measured from the aggregates directly suspended in 0.1 mM R6G and 1 mM BSA (black line), which has much lower intensity than that from the microcapsules.

The aggregate-loaded microcapsules are used as a Raman substrate. As the aggregates have a maximum absorbance at a wavelength of 670 nm as shown in Figure S7b, we select a laser with a wavelength of 633 nm for Raman analysis. We use rhodamine 6G (R6G, Mw 479.02 g/mol) as a target molecule;54 of which hydrodynamic diameter (dH = 1.2 nm) is smaller than the cut-off threshold of 5 nm for microcapsules stressed by ∆C = 100 mOsm/L. Therefore, R6G dissolved in the sample fluids diffuses through the membrane into the aggregates in the core. Raman spectrum measured from the aggregates in the microcapsules shows characteristic peaks of R6G which are located at wavenumbers of 613, 772, 1181,



1315, 1364, 1510, and 1649 cm-1;55 where the intensity of the Raman signal is highest at the central part of aggregates, as shown in Figure S8. This is because the aggregates provide hot spots for SERS. Compared to gold nanoparticles encapsulated in the microcapsules, the aggregates of gold nanoparticles in the microcapsules show 4 times higher intensity of Raman signal when measured from the central part of aggregates, as shown in Figure S9. When measured from the central part of aggregates, the intensity of the peaks decreases along with the concentration of R6G in the sample fluids, as shown in Figure 4c. The concentration and intensity show a linear relation in a log-log scale, as shown in Figure 4d. The limit of detection (LOD) is estimated as 1 nM. In general, metal nanoparticles are prone to contamination by irreversible adsorption of proteins such as albumin.56 Therefore, it is difficult to measure the Raman spectrum of small molecules in the presence of large adhesive proteins. When the aggregate-loaded microcapsules are suspended in the aqueous solution of R6G and FITC-tagged bovine serum albumin (BSA), the microcapsules allow the infusion of R6G, while rejecting FITC-tagged BSA, as shown in Figure 4e. As the hydrodynamic diameter of BSA (dH = 6.78 nm) is larger than the cut-off threshold of 5 nm, BSA is excluded from the microcapsules and the aggregates remain uncontaminated.57 At the same time, R6G with dH = 1.2 nm diffuses through cracks into the core of microcapsules. Therefore, the Raman spectrum of R6G can be measured in the absence of interruption from BSA. When the aqueous solution contains 0.1 mM R6G and 1 mM BSA, Raman spectrum of R6G is selectively measured in high intensity, as denoted by the red curve of Figure 4f. When the aqueous solution contains 0.1 mM R6G only, the intensity is comparable to that from the binary solution of R6G and BSA, as denoted by the blue curve. This indicates that the microcapsules reject BSA and there is negligible interference to the measurement of R6G. When the aggregates of gold nanoparticles are directly suspended in the aqueous solution of 0.1 mM R6G and 1 mM BSA, very low intensity of Raman spectrum is acquired, as denoted with the black curve. This is attributed to the adsorption of BSA on the surface of the aggregates. The microcapsule-type SERS substrate is potentially useful for on-site analysis of complex sample fluids such as biofluids, foods, and cosmetics, as no pretreatment of samples is required.

CONCLUSION Microcapsules with molecular size-selective permeability are promising for various applications. However, there is a lack of methods to produce such microcapsules that are compatible with delicate biological materials and retain dispersion stability of encapsulants while providing high mechanical stability of membranes. Although our approach exerts osmotic stress on living organisms in the core of microcapsules, the stress only lasts for a short time. More importantly, there is no use of chemical etchants to produce pores, which ensures dispersion stability. The membrane is made of entangled polymers also ensuring that it is mechanically stable. Taking advantage of all of these benefits, we successfully design SERS-based molecular sensors using microcapsules with size-selective permeation, which prevents the contamination of sensors through rejection of large adhesive molecules. Our approach is also appealing for encapsulation of cells as it provides a way to isolate cells

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from their surroundings while allowing the controlled exchange of nutrients and metabolites. As a demonstration, we encapsulate yeast cells in the microcapsules and impose a hypotonic stress to make cracks. When the microcapsules are suspended in yeast extract peptone dextrose medium, the yeast cells proliferate within the microcapsules, as shown in Figure S10. In contrast, the microcapsules without any osmotic stress show almost no proliferation of yeast cells as transmembrane transport is prohibited. We believe that the high dispersion stability of encapsulants and biocompatibility of our approach will also provide new opportunities for various applications of the microcapsules with semipermeable membrane beyond the examples demonstrated above.

EXPERIMENTAL SECTION Preparation of a microfluidic device and its operation. The capillary microfluidic device was composed of two tapered cylindrical capillaries assembled in a square capillary, as illustrated in Figure 1. Two cylindrical capillaries (1B100F-6, World Precision Instruments, Inc.) were tapered by a micropipette puller (P97, Sutter Instrument), which were then sanded. One with a 100-µmorifice was treated with octadecyltrimethoxy silane (SigmaAldrich) to render the surfaces hydrophobic and the other with a 180-µm-orifice was treated with 2[methoxy(polyethyleneoxy)propyl] trimethoxy silane (Gelest, Inc.) to render the surfaces hydrophilic. These two cylindrical capillaries were inserted into two openings of a square capillary (OD 1.5 mm, ID 1.05 mm, Atlantic International Technologies, Inc.) to have a tip-to-tip configuration with a separation of 115 µm. A long-tapered cylindrical capillary was inserted into the untapered opening of the hydrophobic capillary. The interstitial opening of the square capillary with the hydrophilic capillary was sealed by an epoxy resin. As the innermost and continuous phases, we used an aqueous solution of 5% w/w PVA (Mw 13,000 – 23,000, SigmaAldrich) and 150 mM NaCl. The middle phase was 10% w/w toluene solution of PLGA (85 mol% DL-lactide: 15 mol% glycolide, Mw 113,000 g/mol, Evonik). The innermost and middle phases were injected into the long-tapered capillary and hydrophobic capillary, whereas the continuous phase was injected through gaps between the hydrophobic capillary and square capillary. The injection was done by syringe pumps (Legato 100, KD Scientific) and typical volumetric flow rates of the innermost, middle, and continuous phases were 400, 200, and 3000 µl/h, respectively. The formation of emulsion drops was observed through inverted optical microscopy (Eclipse TS100, Nikon) with a high-speed camera (Phantom v7.3, Vision Research Inc.). The emulsion drops were collected in a glass dish containing an aqueous solution of 2% w/w PVA and 170 mM NaCl. The single oil drops floated in the collection liquid and were removed by pipetting. The double-emulsion drops were incubated at room temperature for 4 hours. The resulting PLGA microcapsules were observed with optical microscopy. Microcapsules were also inspected with scanning electron microscopy (SEM, S-4800, Hitachi) after fully drying water in a convection oven at 70°C; the microcapsules were washed with distilled water several times before drying and coated with 2-nm-thick Osmium tetroxide after drying for SEM observation. Membrane cross-sections were obtained by randomly cutting the membranes with a razor blade. Crack formation by osmotic pressure difference and analysis of transmembrane transport. PLGA microcapsules were incubated in aqueous solutions with six different osmolarities for 1 day. All solution contains 5% w/w PVA and different concentra-


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tions of NaCl. Osmolarities of all the aqueous solutions were measured by osmometer (Osmomat 030-D, Gonotec) before use. To investigate cut-off threshold of permeation, various fluorescent dyes were dissolved in the surrounding medium, which were sulforhodamine B (Sigma-Aldrich), FITC-tagged dextran with six different molecular weights (Mw 4,000, 10,000, 20,000, 40,000, 70,000, and 150,000 g/mol, Sigma-Aldrich), and RITC-tagged dextran with molecular weight (Mw 10,000 g/mol, SigmaAldrich). After 1 day of incubation, the microcapsules were observed with a confocal laser scanning microscopy (LSM 510, Carl Zeiss). To show size-selective infusion, two dyes with different sizes and fluorescence wavelengths were simultaneously dissolved in the solution. To monitor the temporal change of fluorescence intensity, the microcapsule was periodically observed by a confocal microscope as soon as sulforhodamine B was added in the surrounding solution.

Supporting Information. This material is available free of charge via the Internet at http://pubs.acs.org. OM images of double-emulsion drops and ruptured single-emulsion drops; OM images of PLGA microcapsules at isotonic, hypertonic, hypotonic conditions; SEM images showing dried microcapsules, membrane cross-section, and crack-like features; CLSM image of microcapsules suspended in dye solution; CLSM images of microcapsules showing release of encapsulated dye in various osmotic-pressure conditions; OM image and absorption spectra of dispersions of gold nanoparticles and aggregates; Raman spectra acquired from gold nanoparticles, aggregates, aggregate-loaded microcapsules; OM images of yeast-loaded microcapsules (PDF) Microfluidic production of double-emulsion drops with ultra-thin shell (AVI)

Preparation of microcapsules containing aggregates of gold nanoparticles and Raman analysis. Gold nanoparticles capped by citrate with an average diameter of 20 nm were synthesized by following the protocol previously reported.58 In the aqueous dispersions of 1.4% w/w gold nanoparticles, copper(II) sulfate pen-

AUTHOR INFORMATION

tahydrate (CuSO4 •5H2O) was dissolved in concentrations of 0, 0.5, and 1.0 mM to cause the aggregation of gold nanoparticles. After the formation of aggregates, 5% w/w PVA and 25 mM NaCl were dissolved in the dispersion. The dispersions were observed with optical microscopy and absorbance spectra were measured with a UV-visible spectrometer (Infinite M200 Pro, Tecan). The dispersion was used as the innermost phase and aqueous solution of 5% w/w PVA and 25 mM NaCl was used as the continuous phase to form double-emulsion drops while maintaining the same middle phase. The double-emulsion drops were collected in an aqueous solution of 2% w/w PVA and 40 mM NaCl and incubated at room temperature for 4 hours. The microcapsules were suspended in distilled water for 1 day to impose hypotonic osmotic pressure. The aggregate-loaded microcapsules were suspended in aqueous solutions of R6G (Sigma-Aldrich) at various concentrations in the range of 10-3 M – 10-9 M for 1 day. Raman spectra were acquired from the central part of aggregates in the microcapsules using a dispersive Raman spectrometer (Horiba Jobin Yvon) with a laser wavelength of 633 nm, a power of 6.5 mW, a spot diameter of 1 µm, and acquisition time of 10 s. To study the molecule-selective Raman analysis, 1 mM BSA (Sigma-Aldrich) was co-dissolved in the aqueous solution of 0.1 mM R6G and Raman spectra were measured from the aggregates. To visualize the selective permeation of R6G, the microcapsules suspended in the aqueous solution of R6G and FITC-tagged BSA (Sigma-Aldrich) were observed with a confocal microscopy. For a comparison, aggregates of gold nanoparticles in the absence of microcapsule were also employed.

Author Contributions

Encapsulation of yeast cells. Yeast (Saccharomyces) cells (YSC1, Sigma) were suspended in phosphate-buffered saline (PBS, Thermo Fisher Scientific) solution containing 2% w/w PVA, which was used as the innermost aqueous phase. An aqueous solution of 10% w/w PVA and 100 mM NaCl were used as the continuous phase. The yeast-laden microcapsules were incubated in the aqueous solution of 100 mM NaCl for 2 hours to impose hypotonic osmotic pressure, which were then transferred into a yeast extract peptone dextrose (YEPD, LPS solution) medium. The microcapsules were observed with an optical microscopy after 2 days of incubation at 36°C. For a comparison, the microcapsules were incubated at isotonic condition without osmotic stress and observed after 2 days.

ASSOCIATED CONTENT

Corresponding Author *E-mail: [email protected] (S.-H.K.)

S.L. carried out all experiment; T.Y.L, D.J.K., and B.K helped the characterization; S.-H.K. designed and supervised the research.

Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT This work was supported by the Midcareer Researcher Program (NRF-2017R1A2A2A05001156) and Global Research Laboratory (NRF-2015K1A1A2033054) through the National Research Foundation (NRF) grant funded by the Ministry of Science, ICT and Future Planning (MSIP). We thank Dr. Eugene Hwang for helpful feedback on our manuscript.

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Osmotic-Stress-Mediated Control of Membrane Permeability of

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