Controlling the Stability and Size of Double-Emulsion-Templated Poly

Jun 5, 2012 - The stability and size of poly(lactic-co-glycolic)acid (PLGA)-containing double emulsions and the resulting PLGA microcapsules are contr...
0 downloads 10 Views 3MB Size
Download PDF
Article pubs.acs.org/Langmuir

Controlling the Stability and Size of Double-Emulsion-Templated Poly(lactic-co-glycolic) Acid Microcapsules Fuquan Tu and Daeyeon Lee* Department of Chemical and Biomolecular Engineering, University of Pennsylvania, Philadelphia, Pennsylvania 19104, United States S Supporting Information *

ABSTRACT: The stability and size of poly(lactic-co-glycolic)acid (PLGA)-containing double emulsions and the resulting PLGA microcapsules are controlled by varying the composition of highly monodisperse water-in-oil-in-water (W/O/W) double emulsions. We propose that the basic inner phase of W/O/W double emulsions catalyzes the hydrolysis of PLGA and the ionization of carboxylic acid end groups, which enhances the surface activity of PLGA and facilitates the stabilization of the double emulsions. The size of PLGA-containing double emulsions and that of resulting microcapsules can be readily tuned by osmotic annealing, which depends on the concentration ratio of a solute in the inner and outer phases of double emulsions. The internal volume of PLGA microcapsules can be changed by more than 3 orders of magnitude using this method. This approach also overcomes the difficulty in generating monodisperse double emulsions and microcapsules over a wide range of dimensions using a single microfluidic device. The osmotic annealing method can also be used to concentrate encapsulated species such as colloidal suspensions and biomacromolecules.

■

INTRODUCTION Hollow microcapsules encapsulating active ingredients such as drugs,1−10 genes,11−17 proteins,18−23 and living cells24−34 inside a polymer shell are essential for targeted delivery and sustained release applications.35−39 Hollow microcapsules made of PLGA are ideal systems for numerous biomedical applications because of their excellent biocompatibility and tunable biodegradability.40−42 Conventionally, PLGA microcapsules are fabricated by a technique that involves double emulsification followed by solvent evaporation/extraction.43,44 In this method, an aqueous solution containing active ingredients is emulsified in a PLGA-containing organic solution to form water-in-oil (W/O) emulsions. Subsequently, these W/O emulsion are emulsified in another aqueous solution such as PVA or PVP solution to form water-in-oil-in-water (W/O/W) double emulsions. To remove the organic solvent and form PLGAshelled microcapsules, the W/O/W emulsion is kept either at a reduced pressure or in a large water reservoir to accelerate the removal of the organic solvent. Although great progress in biomedical applications of PLGA microcapsules has been achieved, the conventional method presents a number of challenges that limit the utilization of these microcapsules. For example, the conventional doubleemulsification method has a low encapsulation efficiency, which leads to a significant loss of actives.45,46 In addition, these PLGA microcapsules are very polydisperse in size; therefore, their properties can vary from batch to batch, making it difficult for subsequent characterization and utilization. Furthermore, the shells of these PLGA microcapsules tend to be very porous, © 2012 American Chemical Society

which is thought to be the main reason for the undesirable burst release of encapsulated materials.47 Recent studies have shown that microfluidic techniques provide a versatile means to generate microparticles and hollow microcapsules with high uniformity and encapsulation efficiency.48−55 A microfluidic technique that is particularly useful for the generation of hollow microcapsules relies on the generation of monodisperse water-in-oil-in-water (W/O/W) double emulsions, which are subsequently used to template microcapsule formation. The ability to produce monodisperse droplets also enables the precise characterization and investigation of double emulsions and resulting microcapsules.56 This double-emulsion template method is versatile because it allows the use of different types of materials besides polymers to construct hollow microcapsules.49,50,53,57,58 Another important advantage of the microfluidic approach is that double-emulsion generation becomes a continuous process. This is a significant advantage over other techniques that may be able to provide monodisperse microcapsules. Controlling the size of double emulsions while maintaining high stability and encapsulation efficiency is essential for a variety of applications. Recent studies, however, have shown that it is challenging to control the size of double emulsions and that of resulting microcapsules over a wide range without changing the channel dimensions of the microfluidic device or Received: April 12, 2012 Revised: May 26, 2012 Published: June 5, 2012 9944

dx.doi.org/10.1021/la301498f | Langmuir 2012, 28, 9944−9952

Langmuir

Article

the properties of the fluids, such as viscosity, drastically.59 It was found that a rather small window of flow rates exists, within which monodisperse double emulsions can be consistently generated. Outside this window, single emulsions or double emulsions with multiple inner drops are generated, making it difficult to control the size of double emulsions while maintaining a high encapsulation efficiency. Even if the system is in the stable regime for generating monodisperse double emulsions, the possibility of changing the dimensions of double emulsions (e.g., radius of the inner droplets) by tuning the flow rates was found to be rather limited.59 For example, it is extremely difficult to generate double emulsions that would template microcapsules with inner diameters of 10 and 100 μm using a single microfluidic device, even if the flow rates are changed drastically. A new microfluidic device with appropriate dimensions would have to be fabricated to generate a double emulsion with specific dimensions.56 Therefore, a facile method for controlling the size of monodisperse double emulsions without the need to fabricate a new microfluidic device with different channel dimensions would be highly desirable. The objectives of this study are twofold. We study the stability of PLGA-containing double emulsions and find that the stability of PLGA double emulsions depends on the pH of the inner aqueous phase. We also demonstrate that the size of double emulsions can be controlled within a wide range without utilizing multiple microfluidic devices. We develop a method to control the size of double emulsions and the resulting microcapsules using the osmotic pressure difference between the inner and outer phases of the double emulsions. Although the osmotic pressure has previously been used to vary the size and properties of double emulsions, its use has been limited to highly polydisperse emulsions,60−64 complicating the quantitative analysis of the process. The use of highly monodisperse double emulsions from microfluidics enables the quantitative analysis and prediction of the size of double emulsions and resulting microcapsules. More importantly, this osmotic annealing technique overcomes a major drawback of microfluidic techniques, which do not allow for variation in the size of double emulsions and microcapsules over a wide range using a single microfluidic device. This technique can also be used to concentrate the encapsulated species, which could potentially be useful for encapsulating highly concentrated biomolecular solutions and colloidal suspensions.

■

device, as described previously.65 To generate double emulsions, three different fluid phases are injected into a microfluidic device by three syringe pumps (PHD, Harvard Apparatus) with controlled flow rates. The outer phase is a 2 wt % PVA aqueous solution. The middle phase is 0.9 wt % PLGA in CHCl3. Different salt solutions are used as the inner phase. To generate double emulsions containing a pH indicator dye, Alizarin Yellow R is dissolved in 0.01 and 1 M NaOH solution with a concentration of 100 mg/L. These solutions are used as the inner phase. The middle phase is 0.9 wt % PLGA (50:50) (mol % of lactide/mol % of glycolide) in CHCl3, and the outer phase is a 2 wt % PVA solution. The double emulsions generated using 0.01 M NaOH and 1 M NaOH solutions as the inner phases are collected in 0.01 M NaCl and 1 M NaCl solutions, respectively, to prevent the dilution of the inner phase because of the difference in osmotic pressure between the inner phase and the solution used to collect the double emulsions. These double emulsions are collected in small vials to compare the change in the color of the inner droplets. Stability of PLGA Double Emulsions. PLGA-containing double emulsions encapsulating aqueous solutions of different pH are generated and collected in a Petri dish with deionized water. PLGA with compositions of 50:50, 65:35, and 85:15 are used. The solution pH is tuned by adding NaOH in deionized water. The double emulsions are observed under an upright optical microscope as soon as they are generated. A full set of images for one sample is taken 5 to 10 min after emulsion collection. All of the stable double emulsions become PLGA microcapsules at around 2 h, during which the organic solvent in the middle phase is completely evaporated or removed. The fraction of stable double emulsions is determined by dividing the number of stable double emulsions by the total number of double emulsions collected. Stable double emulsions refer to double emulsions whose inner aqueous droplets remain inside the middle phase until they become PLGA microcapsules. Unstable double emulsions refer to double emulsions whose inner aqueous droplets coalesce with the outer aqueous phase and become single oil-in-water droplets. Unstable double emulsions that become single oil-in-water droplets do not undergo coalescence with each other because they are stabilized by PVA (Figure S1 in Supporting Information). The total number of generated double emulsions is determined by counting the number of stable double emulsions and the number of single emulsions (unstable double emulsions) in optical microscopy images. We analyze over 3200 double emulsions for each data point to obtain the average and standard deviation. Interfacial Tension Measurement. Interfacial tension between the oil phase (i.e., 0.9 wt % PLGA with three compositions in CHCl3) and the aqueous phase (i.e., aqueous solution with different pH's) is determined by pendant drop tensiometry (Attension, Theta). The pH of the aqueous solution is tuned by adding NaOH to the aqueous solution. PLGA Molecular Weight Determination. PLGA (0.9 wt %) in CHCl3 is mixed with 1 M NaOH and PBS solution in volume ratios of 3:0.6 and 3:1, respectively. The mixture is vigorously agitated using a vortex mixer (Fisher Scientific). After vigorous mixing, 2 mL of the oil phase is transferred to another vial. The CHCl3 is removed by evaporation under vacuum. THF (11 mL) is added to the vial to dissolve the solid residue. The vial is placed in a sonicator to help dissolve the PLGA. PLGA in THF is used to determine the molecular weight of PLGA by gel permeation chromatography (Waters) equipped with a refractive index detector. Controlling the Size of Double Emulsions and PLGA Microcapsules Using Osmotic Annealing. Double emulsions are generated with 2 wt % PVA solution and 0.9 wt % PLGA (85:15) as the outer phase and the middle phase, respectively. NaCl solution (0.1 M) with 5 mM Na2CO3 is used as the inner phase to shrink the inner droplet of double emulsions, whereas 0.8 M NaCl is used as the inner phase to swell the inner drops. Generated double emulsions are collected in cuvettes filled with 0.1, 0.2, 0.4, and 0.8 M NaCl solutions. Upon the collection of double emulsions, the cuvettes are capped with stoppers to suppress the evaporation of CHCl3. After the inner droplets reach their equilibrium size, the stoppers are removed to allow

EXPERIMENTAL SECTION

Materials. Deionized water is generated from a Barnstead NANOpure (Thermo Scientific). Ester-terminated PLGA with compositions of 50:50 (intrinsic viscosity (ηi) = 0.41 dL g−1 in hexafluoroisopropanol (HFIP)), 65:35 (ηi = 0.55−0.75 dL g−1 in HFIP), and 85:15 (ηi = 0.66 dL g−1 in CHCl3) are purchased from Durect Corp. A silica nanoparticle suspension (KE-E30) is obtained from Nippon Shokubai, Japan. Poly(vinyl alcohol) (PVA, 87−89% hydrolyzed, average Mw = 13 000−23 000), phosphate-buffered saline (PBS), and calcein are purchased from Sigma-Aldrich. Chloroform (CHCl3), tetrahydrofuran (THF), sodium chloride (NaCl), sodium sulfate (Na2SO4), sodium sulfite (Na2SO3), sodium nitrate (NaNO3), sodium bicarbonate (NaHCO3), sodium carbonate (Na2CO3), sodium hydroxide (NaOH), sodium phosphate monobasic (NaH2PO4), sodium phosphate dibasic (Na2HPO4), potassium chloride (KCl), and potassium phosphate dibasic (K2HPO4) are purchased from Fisher Scientific. Sodium phosphate tribasic (Na3PO4) is purchased from Acros Organics. Alizarin Yellow R was purchased from GFS Chemicals. Double-Emulsion Generation. Water-in-oil-in-water (W/O/W) double emulsions are generated using a glass capillary microfluidic 9945

dx.doi.org/10.1021/la301498f | Langmuir 2012, 28, 9944−9952

Langmuir

Article

of the aqueous phase. We find that salts that do not make the inner phase basic fail to stabilize double emulsions, whereas those that increase the pH of the inner phase are able to stabilize the double emulsions (Figure S1 in the Supporting Information). These results lead us to relate the stability of PLGA-containing double emulsions to the pH of the inner phase rather than the ionic strength. We test our hypothesis by generating PLGA-containing double emulsions with an inner phase of varying pH without extra added salt. For all three types of PLGA used in this study, the stable fraction abruptly increases as the pH of the inner phase reaches 12.67 Double emulsions generated with an inner phase of pH 11 or smaller are unstable (Figures S2 and S3 in the Supporting Information). This result again clearly indicates that a basic environment (i.e., the presence of hydroxide anions) leads to stable double emulsions. It is also important to note that PLGA in the middle phase is necessary to maintain the stability of double emulsions. Double emulsions generated without PLGA in the middle phase, regardless of the innerphase condition, are unstable (Figure S1 in the Supporting Information). Therefore, both a basic inner phase and PLGA are necessary to stabilize the double emulsions. We study the effect of inner-phase pH on the interfacial tension between the aqueous phase and the PLGA-containing oil phase using pendant drop tensiometry. We measure the interfacial tension between chloroform, with and without PLGA, and an aqueous solution as a function of the solution pH. Consistent with the ability of PLGA to stabilize double emulsions, PLGA is found to decrease the interfacial tension between the aqueous and oil phases, as shown in Figure 1.

for the fast evaporation of CHCl3. PLGA microcapsules are formed after the complete evaporation of CHCl3. Concentrating Encapsulated Species Using Osmotic Annealing. NaH2PO4 and Na2HPO4 are added to deionized water in a mole ratio of 22.6:77.4 to make 0.5 mM phosphate-buffered saline (PBS). Then a fluorescent dye, calcein, is added to this 0.5 mM PBS solution to make its concentration 0.1 mg/L. This calcein-containing solution is used as the inner phase to generate double emulsions. Generated double emulsions are collected in either 0.5 mM PBS solution or 0.2 M NaCl solution. Both bright-field and fluorescent images are taken after PLGA microcapsules are formed. To encapsulate silica particles, the original silica nanoparticle suspension (concentration 20 wt %) is centrifuged at 8000 rpm for 5 min. After the removal of supernatant, deionized water is added and the sample is placed in a sonicator for 15 min to redisperse the silica nanoparticles. The process is repeated five times. Subsequently, silica nanoparticles are dried under vacuum. These dried silica nanoparticles are added to a 5 mM Na2CO3 aqueous solution to prepare 0.5 wt % silica nanoparticles. After the nanoparticles are redispersed by sonication, the nanoparticle suspension is used as the inner phase to generate double emulsions with 0.9 wt % PLGA (50:50) in CHCl3 and 2 wt % PVA solution as the middle phase and outer phase, respectively. Generated double emulsions are collected in a 1.0 M NaCl solution and observed under an upright optical microscope.

■

RESULTS AND DISCUSSIONS Stability of PLGA-Containing Double Emulsions. Our previous study has shown that PLGA can stabilize the inner water−oil interface of water-in-oil-in-water (W/O/W) double emulsions without extra surfactants present in the oil phase.66 However, we find that when double emulsions are generated with deionized water (pH 5.5−6.0) as the inner phase such double emulsions are unstable (Figure S1 in Supporting Information). Interestingly, when a PBS solution is used as the inner phase, the inner droplets remain stable inside the oil phase, and hollow PLGA microcapsules can be generated by removing the solvent.66 These results indicate that either the ionic strength and/or the pH of the inner phase plays an important role in controlling the stability of PLGA-containing double emulsions. To gain insight into the stabilization of double emulsion by PLGA, we explore parameters such as the ionic strength and pH of the inner phase as well as the composition of PLGA. We investigate the effect of ionic species in the inner phase. Two types of salts are tested: one that dissociates in water but does not significantly affect the pH of the solution such as NaCl, Na2SO4, NaNO3, and KCl and one that dissociates and forms basic solutions such as Na3PO4, K2HPO4, Na2SO3, and Na2CO3. Table 1 summarizes the effect of each salt on the pH Table 1. Salts Used to Test the Stability of PLGA-Containing Double Emulsions middle phase

inner phase

pH

0.9 wt % PLGA (85:15) in CHCl3

0.05 M K2HPO4 0.05 M Na2SO3 0.005 M Na3PO4 0.05 M Na2CO3

9.00 9.70 11.34 11.29

stable

0.9 wt % PLGA (85:15) in CHCl3

deionized water 0.05 M NaCl 0.05 M KCl 0.05 M Na2SO4 0.05 M NaNO3 0.05 M Na2CO3

5.91 5.81 5.75 6.14 5.94 11.29

unstable

pure CHCl3

Figure 1. Interfacial tension between a PLGA solution in chloroform and an aqueous solution as a function of solution pH. Black squares represents pure chloroform. Red circles, blue triangles, and green inverted triangles represent chloroform solutions with 85:15, 65:35, and 50:50 PLGA, respectively.

stability

Interestingly, the surface activity depends on the composition of PLGA. We believe that the hydrophile−lipophile balance (HLB) of PLGA increases as the fraction of more hydrophilic glycolide is increased in the backbone of PLGA. More importantly, the interfacial tension between the aqueous phase and PLGA-containing chloroform decreases significantly as the pH of the aqueous phase is increased, whereas the 9946

dx.doi.org/10.1021/la301498f | Langmuir 2012, 28, 9944−9952

Langmuir

Article

Table 2. Weight-Averaged Molecular Weight (g/mol) of PLGA after being Placed in Contact with 1 M NaOH Solution and PBS Solution aqueous phase

a

1 M NaOH

PBS

type of PLGA

PLGA (50:50)

PLGA (65:35)

PLGA (85:15)

PLGA (50:50)

0 min 30 min 50 min

63.1 ± 2.9 K 48.6 ± 8.5 K 43.7 ± 2.4 K

50.8 ± 1.7 K 38.7 ± 0.2 K 31.6 ± 0.9 K

68.0 ± 1.7 K 64.5 ± 1.8 K 50.9 ± 1.8 K

62.0 ± 1.8 K 53.5 ± 5.3 K 53.9 ± 5.2 Ka

The molecular weight is measured after the PLGA solution was in contact with the PBS solution for 1 h instead of 50 min.

between the carboxylic acid end groups of PLGA and hydroxide anions in the inner aqueous phase generates ionized carboxylate groups, which are very hydrophilic. These ionized end groups facilitate the anchoring of PLGA at the oil−water interface, which in turn enhances the surface activity of PLGA and its stabilization of the inner droplets of double emulsions. Such a mechanism is consistent with the enhanced double-emulsion stability (Figures S2 and S3 in the Supporting Information) and PLGA surface activity (Figure 1) at a high pH (>12.0). A similar mechanism likely is responsible for the stabilization of W/O/W double emulsions when a PBS solution (pH 7.4) is used as the inner phase. The PBS solution is able to maintain the pH of the inner phase and ionize the carboxylic acid end groups of PLGA, although the hydrolysis of PLGA is not as significant as that of high-pH aqueous solutions (Table 2). We believe that a small number of carboxylic acid groups that were originally present in PLGA could also play an important role in the surface activity of PLGA and the subsequent stabilization of W/O/W double emulsions.80 Controlling the Size of Double Emulsions and PLGA Microcapsules. As described in the Introduction, it is difficult to control the size of double emulsions over a wide range using a single microfluidic device. We overcome this limitation by making use of the osmotic pressure. To tune the size of doubleemulsion-templated microcapsules by what we call osmotic annealing, we separate the formation of microcapsules from double emulsions into two steps, as illustrated in Figure 2. We

change in the interfacial tension between the aqueous phase and pure chloroform as a function of pH is negligible. These results indicate that a high-pH environment enhances the surface activity of PLGA. We further explore the importance of the interaction between basic solutions and PLGA by monitoring the change in the molecular weight of PLGA upon contact with high-pH solutions. To monitor the change in the molecular weight of the polymer, PLGA in CHCl3 is mixed with a 1 M NaOH solution or PBS solution and agitated for different periods of time. Although the oil phase and aqueous phase are immiscible, a significant reduction in the average molecular weight of PLGA is observed with time (Table 2). These results indicate that PLGA undergoes hydrolysis upon contact with basic solutions. Basic solutions have previously been used to render PLGA scaffolds hydrophilic, which have been attributed to the basecatalyzed hydrolysis of ester bonds in PLGA and the formation of carboxylic acid end groups (Figure S4 in the Supporting Information).68−77 In particular, it has been shown that the rate of hydrolysis increases by 5 orders of magnitude when the solution pH increases from 7 to 12.70 We further investigate the importance of the basic condition by monitoring the consumption of hydroxide anions in the inner phase of PLGA-containing double emulsions. To enable this study, we add a pH indicator dye, Alizarin Yellow R, as a probe. Alizarin Yellow R is soluble in water but insoluble in chloroform and thus does not diffuse out of the double emulsions. This dye changes color from dark red to light yellow when the solution pH changes from 12.1 to 10.0 or below. When the dye is dissolved in 0.01 and 1 M NaOH solutions, there is no observable difference in the color of the two NaOH solutions containing Alizarin Yellow R (Figure S5 in the Supporting Information). The color of PLGA double emulsions with a 0.01 M NaOH inner phase changes its color from dark red to light yellow upon collection, whereas PLGA double emulsions with a 1.0 M NaOH inner phase remain red (Figure S5 in the Supporting Information).78 These results indicate that a large number of hydroxide anions in the inner phase are consumed, likely because of the ionization of carboxylic acid end groups of PLGA, leading to the change in the color of PLGA double emulsions and the subsequent pH reduction. It is also likely that monomers of PLGA are released upon hydrolysis.79 Because both monomers, lactic acid and glycolic acid, are highly soluble in water, they could partition into the aqueous phase and contribute to the shift in the pH of the inner droplets. On the basis of these results, we conclude that the ionization of carboxylic acid end groups that form because of the hydrolysis of PLGA chains (Figure S4 in the Supporting Information) is responsible for the stabilization of PLGAcontaining double emulsions. As evidenced by the molecular weight reduction, the base-catalyzed hydrolysis of PLGA provides carboxylic acid end groups. Acid−base reaction

Figure 2. Schematic illustration of tuning the size of double emulsions and microcapsules by osmotic annealing.

generate double emulsions with a salt solution of a given concentration as the inner aqueous phase. Subsequently, we collect double emulsions in a glass cuvette. The glass cuvette is filled with a solution of a different salt concentration. The cuvette is sealed to suppress the removal of the organic solvent in the middle phase. The difference in electrolyte concentrations between the inner droplet and the collection solution induces water to diffuse into and out of the inner droplet, leading to their swelling and shrinkage, respectively. After the inner droplet reaches its equilibrium size, we expose the emulsion suspension to open air to allow the removal of the organic solvent, chloroform. PLGA microcapsules are formed 9947

dx.doi.org/10.1021/la301498f | Langmuir 2012, 28, 9944−9952

Langmuir

Article

Figure 3. (a) PLGA microcapsules formed using osmotic annealing and (b) the diameter evolution of the inner droplets of double emulsions under osmotic annealing. The first, second, and third columns in part a represent the initial double emulsions, double emulsions at equilibrium, and PLGA microcapsules formed, respectively, under different conditions. All of the scale bars in part a are 100 μm. The legend in part b represents the different conditions under which PLGA microcapsules are formed. For example, blue squares with 0.8 M in 0.1 M represent double emulsions generated using a 0.8 M NaCl solution (also containing 5 mM Na2CO3) as the inner phase, collected in 0.1 M NaCl solution.

after the complete removal of the organic solvent as shown in Figure 2. To test the feasibility of this osmotic annealing, we generate eight samples of double emulsions with the same initial dimensions as shown in Figure 3. On the basis of the results from the first part of this study, 5 mM Na2CO3 is added to the inner phase to enhance the stability of PLGA-containing double emulsions whereas NaCl in the inner phase is used to provide an osmotic pressure difference for osmotic annealing. When the concentration of NaCl in the inner droplets is higher than that in the collection solution, the inner droplets swell. In contrast, the inner droplets shrink if the situation is the opposite. These swelling and shrinking behaviors can be understood by considering the difference in osmolarity (i.e., chemical potential of water) of the inner droplet and the collection solution. When the inner phase is more concentrated, water in the collection solution has a higher chemical potential than that of the inner phase; therefore, water diffuses into the inner droplets from the collection solution through the middle phase. This transport of water makes the inner droplets swell. The inner droplets reach their equilibrium diameter within 2 h of osmotic annealing, which can be inferred from the plateau of the diameter evolution curves in Figure 3b. PLGA microcapsules ranging from 80 to ∼300 μm in diameter are formed from one double emulsion (initial inner and outer diameters were ∼150 and 250 μm, respectively) using osmotic annealing as shown in Figure 3a. The equilibrium diameter of the inner droplets of double emulsions is determined by the balance of osmotic pressure due to the concentration difference between the inner droplet and the collection solution and the Laplace pressure due to interfacial tension at the two oil−water interfaces. To derive the relationship between the salt concentration ratio and the equilibrium diameter of the inner droplet, we assume that there is negligible transport of the ionic species across the middle phase, which is reasonable because the solubility of the ions in chloroform is extremely low. The Laplace pressure associated with the two oil−water interfaces is

P=

2γim rin

+

2γom rout

(1)

where γim is the interfacial tension between the inner and middle phases and γom represents the interfacial tension between the outer and middle phases. rin and rout represent the radii of the inner and outer droplets, respectively. The osmotic pressure difference between the inner and outer phases can be represented by van’t Hoff’s law:81

Π = 2(C in − Cout)RT

(2)

Cin and Cout are the concentrations of NaCl in the inner droplets and the collection solution at equilibrium (Figure S6 in the Supporting Information). R is the gas constant, and T is the absolute temperature. The equilibrium diameter of the inner droplets is determined by equating eqs 1 and 2 as shown in eq 3: 2(C in − Cout)RT =

2γim rin − equ

+

2γom rout − equ

(3)

where rin−equ and rout−equ are the radii of the inner and outer droplets at equilibrium, respectively. Neglecting the transport of the ionic species, the concentration of NaCl in the inner droplet at equilibrium is related to its original value by eq 4, which implies the conservation of ionic species in the inner droplets: ⎛ r 0 ⎞3 C in = C in0⎜⎜ in ⎟⎟ ⎝ rin − equ ⎠

(4)

Substituting eq 4 into eq 3, we obtain ⎛ r 0 ⎞3 γom ⎞ 1 ⎛⎜ γim in ⎟ ⎟ − Cout = + ⎟ ⎜ RT ⎝ rin − equ rout − equ ⎟⎠ ⎝ rin − equ ⎠

C in0⎜⎜

(5)

Solving eq 5, the ratio of rin−equ to r0in can be expressed as 9948

dx.doi.org/10.1021/la301498f | Langmuir 2012, 28, 9944−9952

Langmuir

rin − equ rin0

⎛ ⎜ =⎜ ⎜⎜ C + ⎝ out

Article

C in0

1 ⎛ γim ⎜ RT ⎝ rin − equ

⎞1/3 ⎟ ⎟ ⎞ γ + r om ⎟ ⎟⎟ out − equ ⎠ ⎠

middle phase, the volume of the middle phase, the density of PLGA, and the equilibrium inner diameter of PLGA microcapsules (eqs S1 and S2 in the Supporting Information). We can make further use of osmotic annealing to concentrate encapsulated species in double emulsions and microcapsules. We use a 0.5 mM phosphate buffer solution (PBS) with 0.1 mg/L calcein in the inner phase as a probe dye to generate double emulsions. When these double emulsions are collected in 0.5 mM PBS and converted to PLGA microcapsules, very little fluorescence intensity is detected because of the low concentration of calcein (Figure 5a). In contrast, when these double emulsions are collected in 0.2 M NaCl solution, water inside the inner droplets diffuses out through the oil phase because of the difference in osmolarity between the inner and outer phases, whereas calcein remains encapsulated in the inner droplets. Consequently, the concentration of calcein and, in turn, the fluorescence intensity of the inner phase increase substantially. The increase in fluorescence intensity is quite drastic as seen in Figure 5b,c, which shows a line scan of fluorescence in the microcapsules generated with and without osmotic annealing. It is worth noting that in our swelling experiment the volume of the inner droplets is increased by a factor of 8 (Figure 3b) whereas in the shrinking experiment the volume of the inner droplets is decreased by a factor of ∼270 (Figure 5a,b); therefore, our results indicate that the volume of the inner droplets and, in turn, the concentration of the encapsulated solute can be tuned by at least 3 orders of magnitude using the osmotic annealing method. This concentration method using osmotic annealing is especially useful in encapsulating highly concentrated solid suspensions such as colloidal particles. Typically, microfluidic channels clog rapidly when highly concentrated solutions of suspensions are used.82,83 We can readily encapsulate highly concentrated particle suspension by annealing double emulsions containing a low initial concentration of particles in a high salt concentration solution. We demonstrate this by essentially squeezing out all of the water from encapsulated particle suspensions, which leaves an agglomerate of particles in the core of oil droplets as shown in Figure 5d.

(6)

We find that (1/(RT))((γim)/(rin−equ) + ((γom)/(rout−equ))) is approximately or less than 1% of C0in and Cout (Supporting Information). Thus, we can simplify eq 6 by neglecting the effect of interfacial tension on the equilibrium diameter of the inner droplet. Thus, we obtain a very simple relationship between the initial and the equilibrium radii of the inner droplet as shown in eq 7: rin − equ rin0

⎛ C 0 ⎞1/3 = ⎜ in ⎟ ⎝ Cout ⎠

(7)

The diameter of the inner droplets after osmotic annealing can be accurately predicted by the theoretical model as represented by eq 7. The relationship between the one-third power of the concentration ratio ((C0in/Cout)1/3) and the radii ratio (rin−equ/r0in) for eight different concentration ratios is shown in Figure 4. Note that the diameter of the inner droplet

■

CONCLUSIONS We have demonstrated that the stability and size of PLGAcontaining double emulsions and resulting PLGA microcapsules can be controlled by varying the composition of each phase of the double emulsions. In particular, it is important to adjust the pH of the inner phase to enable the stabilization of inner droplets of PLGA-containing double emulsions. The hydrolysis of PLGA and, more importantly, the ionization of carboxylic acid end groups enhance the surface activity of PLGA at the oil−water interface and, in turn, the stabilization of PLGA-containing double emulsions. We have also shown that the size of double emulsions can be precisely tuned by changing the concentration ratio of a solute in the inner and outer phases of double emulsions. It is possible to change the volume of the internal compartment of PLGA microcapsules by more than 3 orders of magnitude, illustrating the effectiveness and reliability of this new method in changing the size of microcapsules and, in turn, the concentration of the encapsulated species. This approach overcomes the difficulty in generating monodisperse double emulsions and microcapsules over a wide range of dimensions using a single microfluidic device. We believe that the osmotic annealing method will be useful in generating drug-delivery systems encapsulating a high

Figure 4. Ratio of initial and equilibrium diameters of inner droplets as a function of the NaCl concentration ratio of the inner and collection solutions. The dotted line represents the prediction based on eq 7. The legend uses the same notation as in Figure 3.

measured from optical microscopy is different from its actual diameter because of the difference in the refractive indices of water and organic solvents. All data for the diameter of the inner droplets are corrected using a method provided elsewhere.59 It can be clearly seen that the experimental data from Figure 3 all lie on the line determined by eq 7, indicating that the experimental results agree well with our hypothesis and prediction. Most importantly, these results strongly support the fact that we can readily tune the size of the microcapsules and that of the inner droplets of double emulsions from a single device under one set of double-emulsion formation conditions by simply changing the ratio of the solute concentrations of the inner and collection solutions. The thickness of the microcapsule shell is an important factor that determines the release of encapsulated species. The average shell thickness can be determined on the basis of the concentration of PLGA in the 9949

dx.doi.org/10.1021/la301498f | Langmuir 2012, 28, 9944−9952

Langmuir

Article

Figure 5. Concentrating encapsulated species using osmotic annealing. (a, b) Bright field and fluorescent images of the PLGA microcapsules without and with osmotic annealing. (c) Fluorescence intensity of encapsulated species along the white lines in a and b. (d) Optical microscopy images showing the shrinkage of an encapsulated silica suspension to an agglomerate of silica particles under osmotic annealing. Double emulsions are generated with 0.5 wt % silica particle suspensions in 5 mM Na2CO3 as the inner phase and collected in 1.0 M NaCl solution.

concentration of biomacromolecules such as proteins84 and also for generating capsules encapsulating a high concentration of solid suspensions while avoiding the clogging of microfluidic devices.

■

doxorubicin loading and in vitro and in vivo studies. Nanomed: Nanotechnol, Biol., Med 2007, 3, 63−74. (3) Liu, X.; Gao, C.; Shen, J.; Möhwald, H. Multilayer microcapsules as anti-cancer drug delivery vehicle: deposition, sustained release, and in vitro bioactivity. Macromol. Biosci. 2005, 5, 1209−1219. (4) An, Z.; Lu, G.; Möhwald, H.; Li, J. Self-assembly of human serum albumin (HSA) and l-α-dimyristoylphosphatidic acid (DMPA) microcapsules for controlled drug release. Chem.Eur. J. 2004, 10, 5848−5852. (5) Fundueanu, G.; Constantin, M.; Esposito, E.; Cortesi, R.; Nastruzzi, C.; Menegatti, E. Cellulose acetate butyrate microcapsules containing dextran ion-exchange resins as self-propelled drug release system. Biomaterials 2005, 26, 4337−4347. (6) Lu, L.; Huang, K. Preparation of poly(propylene-co-γbutyrolactone carbonate) and release profiles of drug-loaded microcapsules. J. Polym. Sci., Part A: Polym. Chem. 2005, 43, 2468−2475. (7) Wang, S.-B.; Xu, F.-H.; He, H.-S.; Weng, L.-J. Novel Alginatepoly(L-histidine) microcapsules as drug carriers: in vitro protein release and short term stability. Macromol. Biosci. 2005, 5, 408−414. (8) Huang, S.-L.; MacDonald, R. C. Acoustically active liposomes for drug encapsulation and ultrasound-triggered release. Biochim. Biophys. Acta, Biomembr. 2004, 1665, 134−141. (9) Wang, C.; Ye, W.; Zheng, Y.; Liu, X.; Tong, Z. Fabrication of drug-loaded biodegradable microcapsules for controlled release by combination of solvent evaporation and layer-by-layer self-assembly. Int. J. Pharm. 2007, 338, 165−173. (10) Song, W.; He, Q.; Mohwald, H.; Yang, Y.; Li, J. Smart polyelectrolyte microcapsules as carriers for water-soluble small molecular drug. J. Controlled Release 2009, 139, 160−166. (11) Ando, S.; Putnam, D.; Pack, D. W.; Langer, R. PLGA microspheres containing plasmid DNA: preservation of supercoiled DNA via cryopreparation and carbohydrate stabilization. J. Pharm. Sci. 1999, 88, 126−130. (12) Wang, D.; Robinson, D. R.; Kwon, G. S.; Samuel, J. Encapsulation of plasmid DNA in biodegradable poly(d,l-lactic-coglycolic acid) microspheres as a novel approach for immunogene delivery. J. Controlled Release 1999, 57, 9−18. (13) Capan, Y.; Woo, B. H.; Gebrekidan, S.; Ahmed, S.; DeLuca, P. P. Preparation and characterization of poly(D,L-lactide-co-glycolide)

ASSOCIATED CONTENT

* Supporting Information S

Images of stable and unstable double emulsions, an estimation of the interfacial tension term in eq 6, and equations for estimating the average shell thickness. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The osmotic annealing and stability study was supported by the PENN MRSEC through the National Science Foundation DMR-1120901 and an NSF CAREER Award (DMR-1055594), and the microfluidic generation of double emulsions was supported by the Biomolecular Materials program at the DOE (DE-FG02-11ER46810). D.L. also acknowledges the support of Korean−American Scientists and Engineers Association (KSEA). We thank Ryan Wade (University of Pennsylvania) for his assistance with the gel permeation chromatography of PLGA.

■

REFERENCES

(1) Waeckerle-Men, Y.; Groettrup, M. PLGA microspheres for improved antigen delivery to dendritic cells as cellular vaccines. Adv. Drug Delivery Rev. 2005, 57, 475−482. (2) Zhao, Q.; Han, B.; Wang, Z.; Gao, C.; Peng, C.; Shen, J. Hollow chitosan-alginate multilayer microcapsules as drug delivery vehicle: 9950

dx.doi.org/10.1021/la301498f | Langmuir 2012, 28, 9944−9952

Langmuir

Article

by radical-free gelation in droplet microfluidics. J. Am. Chem. Soc. 2012, 134, 4983−4989. (35) Langer, R. Drug delivery and targeting. Nature 1998, 392, 5−10. (36) Sukhorukov, G. B.; Rogach, A. L.; Garstka, M.; Springer, S.; Parak, W. J.; Muñoz-Javier, A.; Kreft, O.; Skirtach, A. G.; Susha, A. S.; Ramaye, Y.; Palankar, R.; Winterhalter, M. Multifunctionalized polymer microcapsules: novel tools for biological and pharmacological applications. Small 2007, 3, 944−955. (37) Orive, G.; Hernandez, R. M.; Gascon, A. R.; Calafiore, R.; Chang, T. M. S.; Vos, P. D.; Hortelano, G.; Hunkeler, D.; Lacik, I.; Shapiro, A. M. J.; Pedraz, J. L. Cell encapsulation: promise and progress. Nat. Med. 2003, 9, 104−107. (38) Putnam, D. Polymers for gene delivery across length scales. Nat. Mater. 2006, 5, 439−451. (39) Luo, D.; Saltzman, W. M. Synthetic DNA delivery systems. Nat. Biotechnol. 2000, 18, 33−37. (40) Anderson, J. M.; Shive, M. S. Biodegradation and biocompatibility of PLA and PLGA microspheres. Adv. Drug Delivery Rev. 1997, 28, 5−24. (41) Bala, I.; Hariharan, S.; Kumar, M. N. PLGA nanoparticles in drug delivery: the state of the art. Crit. Rev. Ther. Drug Carrier Syst. 2004, 21, 387−422. (42) Athanasiou, K. A.; Niederauer, G. G.; Agrawal, C. M. Sterilization, toxicity, biocompatibility and clinical applications of polylactic acid/ polyglycolic acid copolymers. Biomaterials 1996, 17, 93−102. (43) Freitas, S.; Merkle, H. P.; Gander, B. Microencapsulation by solvent extraction/evaporation: reviewing the state of the art of microsphere preparation process technology. J. Controlled Release 2005, 102, 313−332. (44) Rajeev, A, J. The manufacturing techniques of various drug loaded biodegradable poly(lactide-co-glycolide) (PLGA) devices. Biomaterials 2000, 21, 2475−2490. (45) Nihant, N.; Schugens, C.; Grandfils, C.; Jerome, R.; Teyssie, P. Polylactide microparticles prepared by double emulsion-evaporation: II. Effect of the poly(lactide-co-glycolide) composition on the stability of the primary and secondary emulsions. J. Colloid Interface Sci. 1995, 173, 55−65. (46) Yeo, Y.; Park, K. Control of encapsulation efficiency and initial burst in polymeric microparticle systems. Arch. Pharm. Res. 2004, 27, 1−12. (47) Wang, H. T.; Schmitt, E.; Flanagan, D. R.; Linhardt, R. J. Influence of formulation methods on the in vitro controlled release of protein from poly (ester) microspheres. J. Controlled Release 1991, 17, 23−31. (48) Xu, S.; Nie, Z.; Seo, M.; Lewis, P.; Kumacheva, E.; Stone, H. A.; Garstecki, P.; Weibel, D. B.; Gitlin, I.; Whitesides, G. M. Generation of monodisperse particles by using microfluidics: control over size, shape, and composition. Angew. Chem. 2005, 117, 734−738. (49) Shum, H. C.; Kim, J.-W.; Weitz, D. A. Microfluidic fabrication of monodisperse biocompatible and biodegradable polymersomes with controlled permeability. J. Am. Chem. Soc. 2008, 130, 9543−9549. (50) Shum, H. C.; Lee, D.; Yoon, I.; Kodger, T.; Weitz, D. A. Double emulsion templated monodisperse phospholipid vesicles. Langmuir 2008, 24, 7651−7653. (51) Choi, S.-W.; Zhang, Y.; Xia, Y. Fabrication of microbeads with a controllable hollow interior and porous wall using a capillary fluidic device. Adv. Funct. Mater. 2009, 19, 2943−2949. (52) Stachowiak, J. C.; Richmond, D. L.; Li, T. H.; Liu, A. P.; Parekh, S. H.; Fletcher, D. A. Unilamellar vesicle formation and encapsulation by microfluidic jetting. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 4697− 4702. (53) Kim, J.-W.; Utada, A. S.; Fernández-Nieves, A.; Hu, Z.; Weitz, D. A. Fabrication of monodisperse gel shells and functional microgels in microfluidic devices. Angew. Chem. 2007, 119, 1851−1854. (54) Lee, M. H.; Lee, D. Elastic instability of polymer-shelled bubbles formed from air-in-oil-in-water compound bubbles. Soft Matter 2010, 6, 4326−4330.

microspheres for controlled release of poly(L-lysine) complexed plasmid DNA. Pharm. Res. 1999, 16, 509−513. (14) Luo, D.; Woodrow-Mumford, K.; Belcheva, N.; Saltzman, W. M. Controlled DNA delivery systems. Pharm. Res. 1999, 16, 1300−1308. (15) Wang, C.; Ge, Q.; Ting, D.; Nguyen, D.; Shen, H.-R.; Chen, J.; Eisen, H. N.; Heller, J.; Langer, R.; Putnam, D. Molecularly engineered poly(ortho ester) microspheres for enhanced delivery of DNA vaccines. Nat. Mater. 2004, 3, 190−196. (16) Walter, E.; Moelling, K.; Pavlovic, J.; Merkle, H. P. Microencapsulation of DNA using poly(dl-lactide-co-glycolide): stability issues and release characteristics. J. Controlled Release 1999, 61, 361−374. (17) Walter, E.; Dreher, D.; Kok, M.; Thiele, L.; Kiama, S. G.; Gehr, P.; Merkle, H. P. Hydrophilic poly(dl-lactide-co-glycolide) microspheres for the delivery of DNA to human-derived macrophages and dendritic cells. J. Controlled Release 2001, 76, 149−168. (18) Yang, Y.-Y.; Chia, H.-H.; Chung, T.-S. Effect of preparation temperature on the characteristics and release profiles of PLGA microspheres containing protein fabricated by double-emulsion solvent extraction/evaporation method. J. Controlled Release 2000, 69, 81−96. (19) Christian, D. A.; Cai, S.; Bowen, D. M.; Kim, Y.; Pajerowski, J. D.; Discher, D. E. Polymersome carriers: from self-assembly to siRNA and protein therapeutics. Eur. J. Pharm. Biopharm. 2009, 71, 463−474. (20) Maulding, H. V. Prolonged delivery of peptides by microcapsules. J. Controlled Release 1987, 6, 167−176. (21) Elvassore, N.; Bertucco, A.; Caliceti, P. Production of proteinloaded polymeric microcapsules by compressed CO2 in a mixed solvent. Ind. Eng. Chem. Res. 2001, 40, 795−800. (22) Balabushevitch, N. G.; Sukhorukov, G. B.; Moroz, N. A.; Volodkin, D. V.; Larionova, N. I.; Donath, E.; Mohwald, H. Encapsulation of proteins by layer-by-layer adsorption of polyelectrolytes onto protein aggregates: Factors regulating the protein release. Biotechnol. Bioeng. 2001, 76, 207−213. (23) Caruso, F.; Trau, D.; Mohwald, H.; Renneberg, R. Enzyme encapsulation in layer-by-layer engineered polymer multilayer capsules. Langmuir 2000, 16, 1485−1488. (24) Tan, W. H.; Takeuchi, S. Monodisperse alginate hydrogel microbeads for cell encapsulation. Adv. Mater. 2007, 19, 2696−2701. (25) Chia, S. M.; Wan, A. C. A.; Quek, C. H.; Mao, H. Q.; Xu, X.; Shen, L.; Ng, M. L.; Leong, K. W.; Yu, H. Multi-layered microcapsules for cell encapsulation. Biomaterials 2002, 23, 849−856. (26) Peirone, M.; Ross, C. J. D.; Hortelano, G.; Brash, J. L.; Chang, P. L. Encapsulation of various recombinant mammalian cell types in different alginate microcapsules. J. Biomed. Mater. Res. 1998, 42, 587− 596. (27) Blasi, P.; Giovagnoli, S.; Schoubben, A.; Ricci, M.; Rossi, C.; Luca, G.; Basta, G.; Calafiore, R. Preparation and in vitro and in vivo characterization of composite microcapsules for cell encapsulation. Int. J. Pharm. 2006, 324, 27−36. (28) Matthew, H. W.; Salley, S. O.; Peterson, W. D.; Klein, M. D. Complex coacervate microcapsules for mammalian cell culture and artificial organ development. Biotechnol. Prog. 1993, 9, 510−519. (29) Chandy, T.; Mooradian, D. L.; Rao, G. H. R. Evaluation of modified alginate-chitosan-polyethylene glycol microcapsules for cell encapsulation. Artif. Organs 1999, 23, 894−903. (30) Keen, P. H. R.; Slater, N. K. H.; Routh, A. F. Encapsulation of yeast cells in colloidosomes. Langmuir 2012, 28, 1169−1174. (31) Trongsatitkul, T.; Budhlall, B. M. Multicore-shell PNIPAm-coPEGMa microcapsules for cell encapsulation. Langmuir 2012, 27, 13468−13480. (32) Hamad, S. A.; Dyab, A. F. K.; Stoyanov, S. D.; Paunov, V. N. Encapsulation of living cells into sporopollenin microcapsules. J. Mater. Chem. 2012, 21, 18018−18023. (33) Mazumder, M. A. J.; Burke, N. A. D.; Shen, F.; Potter, M. A.; Stover, H. D. H. Core-cross-linked alginate microcapsules for cell encapsulation. Biomacromolecules 2009, 10, 1365−1373. (34) Rossow, T.; Heyman, J. A.; Ehrlicher, A. J.; Langhoff, A.; Weitz, D. A.; Haag, R.; Seiffert, S. Controlled synthesis of cell-laden microgels 9951

dx.doi.org/10.1021/la301498f | Langmuir 2012, 28, 9944−9952

Langmuir

Article

(55) Lee, M. H.; Prasad, V.; Lee, D. Microfluidic fabrication of stable nanoparticle-shelled bubbles. Langmuir 2009, 26, 2227−2230. (56) Utada, A. S.; Lorenceau, E.; Link, D. R.; Kaplan, P. D.; Stone, H. A.; Weitz, D. A. Monodisperse double emulsions generated from a microcapillary device. Science 2005, 308, 537−541. (57) Kamat, N. P.; Lee, M. H.; Lee, D.; Hammer, D. A. Micropipette aspiration of double emulsion-templated polymersomes. Soft Matter 2012, 7, 9863−9866. (58) Lee, D.; Weitz, D. A. Double emulsion-templated nanoparticle colloidosomes with selective permeability. Adv. Mater. 2008, 20, 3498−3503. (59) Erb, R. M.; Obrist, D.; Chen, P. W.; Studer, J.; Studart, A. R. Predicting sizes of droplets made by microfluidic flow-induced dripping. Soft Matter 2011, 7, 8757−8761. (60) Mezzenga, R.; Folmer, B. M.; Hughes, E. Design of double emulsions by osmotic pressure tailoring. Langmuir 2004, 20, 3574− 3582. (61) Herrmann, J.; Bodmeier, R. The effect of particle microstructure on the somatostatin release from poly(lactide) microspheres prepared by a W/O/W solvent evaporation method. J. Controlled Release 1995, 36, 63−71. (62) K. F. Pistel, T. K. Effects of salt addition on the microencapsulation of proteins using W/O/W double emulsion technique. J. Microencapsulation 2000, 17, 467−483. (63) Han, K.; Lee, K.-D.; Gao, Z.-G.; Park, J.-S. Preparation and evaluation of poly(l-lactic acid) microspheres containing rhEGF for chronic gastric ulcer healing. J. Controlled Release 2001, 75, 259−269. (64) Jiang, G.; Thanoo, B. C.; DeLuca, P. P. Effect of osmotic pressure in the solvent extraction phase on BSA release profile from PLGA microspheres. Pharm. Dev. Technol. 2002, 7, 391−399. (65) Shah, R. K.; Shum, H. C.; Rowat, A. C.; Lee, D.; Agresti, J. J.; Utada, A. S.; Chu, L.-Y.; Kim, J.-W.; Fernandez-Nieves, A.; Martinez, C. J.; Weitz, D. A. Designer emulsions using microfluidics. Mater. Today 2008, 11, 18−27. (66) Lee, M. H.; Hribar, K. C.; Brugarolas, T.; Kamat, N. P.; Burdick, J. A.; Lee, D. Harnessing interfacial phenomena to program the release properties of hollow microcapsules. Adv. Funct. Mater. 2012, 22, 131− 138. (67) Although the effect of the PLGA composition is not overwhelming, 50:50 PLGA seems to stabilize double emulsions most effectively. At pH 11.0, the fraction of stable double emulsions containing 50:50 PLGA is greater than those containing 65:35 and 85:15 PLGA. (68) Miller, D. C.; Thapa, A.; Haberstroh, K. M.; Webster, T. J. Endothelial and vascular smooth muscle cell function on poly(lacticco-glycolic acid) with nano-structured surface features. Biomaterials 2004, 25, 53−61. (69) Gao, J.; Niklason, L.; Langer, R. Surface hydrolysis of poly(glycolic acid) meshes increases the seeding density of vascular smooth muscle cells. J. Biomed. Mater. Res. 1998, 42, 417−424. (70) Croll, T. I.; O’Connor, A. J.; Stevens, G. W.; Cooper-White, J. J. Controllable surface modification of poly(lactic-co-glycolic acid) (PLGA) by hydrolysis or aminolysis I: physical, chemical, and theoretical aspects. Biomacromolecules 2004, 5, 463−473. (71) Park, G. E.; Pattison, M. A.; Park, K.; Webster, T. J. Accelerated chondrocyte functions on NaOH-treated PLGA scaffolds. Biomaterials 2005, 26, 3075−3082. (72) Smith, L. L.; Niziolek, P. J.; Haberstroh, K. M.; Nauman, E. A.; Webster, T. J. Decreased fibroblast and increased osteoblast adhesion on nanostructured NaOH-etched PLGA scaffolds. Int. J. Nanomed. 2007, 2, 383−388. (73) Nam, Y. S.; Yoon, J. J.; Lee, J. G.; Park, T. G. Adhesion behaviours of hepatocytes cultured onto biodegradable polymer surface modified by alkali hydrolysis process. J. Biomater. Sci., Polym. Ed. 1999, 10, 1145−1158. (74) Khang, G.; Choee, J.-H.; Rhee, J. M.; Lee, H. B. Interaction of different types of cells on physicochemically treated poly(L-lactide-coglycolide) surfaces. J. Appl. Polym. Sci. 2002, 85, 1253−1262.

(75) Pattison, M. A.; Wurster, S.; Webster, T. J.; Haberstroh, K. M. Three-dimensional, nano-structured PLGA scaffolds for bladder tissue replacement applications. Biomaterials 2005, 26, 2491−2500. (76) Thapa, A.; Miller, D. C.; Webster, T. J.; Haberstroh, K. M. Nano-structured polymers enhance bladder smooth muscle cell function. Biomaterials 2003, 24, 2915−2926. (77) Thapa, A.; Webster, T. J.; Haberstroh, K. M. Polymers with nano-dimensional surface features enhance bladder smooth muscle cell adhesion. J. Biomed. Mater. Res., Part A 2003, 67, 1374−1383. (78) One molar (1 M) NaOH solution is not suitable for the encapsulation of active ingredients. This basic solution, however, is used to study the effect of the inner solution basicity on the hydrolysis of PLGA in the oil phase and the ionization of carboxylic acid groups. (79) Giunchedi, P.; Conti, B.; Scalia, S.; Conte, U. In vitro degradation study of polyester microspheres by a new HPLC method for monomer release determination. J. Controlled Release 1998, 56, 53−62. (80) Although ester-terminated PLGA is used, PLGA may undergo slow hydrolysis during storage at low temperature (= −17 °C). PLGA has been shown to undergo hydrolysis at 0 °C.70 (81) Atkins, P.; de Paula, J. Physical Chemistry, 7th ed.; W. H. Freeman and Company: New York, 2001; p 178. (82) Wyss, H. M.; Blair, D. L.; Morris, J. F.; Stone, H. A.; Weitz, D. A. Mechanism for clogging of microchannels. Phys. Rev. E 2006, 74, 061402. (83) Kanai, T.; Lee, D.; Shum, H. C.; Weitz, D. A. Fabrication of tunable spherical colloidal crystals immobilized in soft hydrogels. Small 2010, 6, 807−810. (84) Shire, S. J.; Shahrokh, Z.; Liu, J. Challenges in the development of high protein concentration formulations. J. Pharm. Sci. 2004, 93, 1390−1402.

9952

dx.doi.org/10.1021/la301498f | Langmuir 2012, 28, 9944−9952