Improved Protein Conjugation with Uniform, Macroporous Poly

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Improved Protein Conjugation with Uniform, Macroporous Poly(acrylamide-co-acrylic acid) Hydrogel Microspheres via EDC/NHS Chemistry Eric Y. Liu, Sukwon Jung, and Hyunmin Yi* Department of Chemical and Biological Engineering, Tufts University, Medford, Massachusetts 02155, United States S Supporting Information *

ABSTRACT: We demonstrate a robust and tunable micromolding method to fabricate chemically functional poly(acrylamide-co-acrylic acid) (p(AAm-co-AA)) hydrogel microspheres with uniform dimensions and controlled porous network structures for rapid biomacromolecular conjugation. Specifically, p(AAm-co-AA) microspheres with abundant carboxylate functional groups are fabricated via surfacetension-induced droplet formation in patterned poly(dimethylsiloxane) molds and photoinduced radical polymerization. To demonstrate the chemical functionality, we enlisted rapid EDC/NHS (1-ethyl-3-(3-(dimethylamino)propyl)carbodiimide (EDC) and N-hydroxysuccinimide (NHS)) chemistry for fluorescent labeling of the microspheres with small-molecule dye fluorescein glycine amide. Epifluorescence imaging results illustrate the uniform incorporation of carboxylate groups within the microspheres and rapid conjugation kinetics. Furthermore, protein conjugation results using red fluorescent protein R-phycoerythrin demonstrate the highly porous nature of the microspheres as well as the utility of the microspheres and the EDC/NHS scheme for facile biomacromolecular conjugation. Combined, these results illustrate the significant potential for our fabrication−conjugation strategy in the development of biofunctionalized polymeric hydrogel microparticles toward rapid biosensing, bioprocess monitoring, and biodiagnostics.

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INTRODUCTION Functionalized polymeric hydrogel microparticles have recently drawn considerable attention for a wide range of applications including medical diagnostics,1,2 biosensing,3,4 tissue engineering,5,6 and controlled delivery of therapeutic agents (e.g., proteins, cells, and synthetic drugs)7,8 as a result of their highly tunable nature,9,10 solution-like kinetics,11 potential for substantially higher probe and target binding capacity over surface-based planar arrays,12 nonfouling properties5,9 for selective sensing, and biocompatibility.9 As such, numerous advances in the fabrication of these hydrogel microparticles have been made over traditional fabrication methods such as dispersion or emulsion polymerization.10 For example, microfluidics-based stop-flow lithography methods impart a high multiplexing ability through the fabrication of graphically encoded microparticles.1,11 Droplet-based microfluidics and other microfluidic techniques offer a rapid fabrication of multicompartmental microparticles with complex three-dimensional (3D) structures.13−15 Meanwhile, soft-lithography-based imprinting16,17 and micromolding-based18−20 techniques offer excellent control of particle uniformity, morphology, and size without the need for complex and expensive equipment. However, despite such advances, there still exist critical challenges in the current fabrication techniques for hydrogel microparticle systems. For example, microfluidic systems often require complex devices, surfactants, and high-intensity ultra© XXXX American Chemical Society

violet (UV) light conditions that can limit the control of microparticle fabrication and compromise the functionality of fabricated microparticles.1,11,21 Meanwhile, soft-lithographybased techniques are challenged with the persistent formation of a residual flash layer connecting potential particles during the fabrication process.21 In the meantime, microparticles containing carboxylate functionalities hold the potential for simple and rapid protein conjugation via widely used rapid 1-ethyl-3-(3(dimethylamino)propyl)carbodiimide (EDC)-N-hydroxysuccinimide (NHS) chemistry.22,23 However, existing studies lack precise control of the dimensions and mesh size of carboxylatecontaining microspheres24−26 or neglect to utilize optimal reaction conditions of EDC/NHS in polymeric platforms.7,27−29 Although other groups have previously developed macroporous carboxylate-containing hydrogels, robust carboxylate-functional hydrogel microparticle formats have not yet been realized.30,31 Therefore, there exists a critical need for facile fabrication methods to produce chemically functional polymeric hydrogel microparticle platforms with controlled dimensions and macroporous network structures for biomacromolecular conjugation and biosensing applications. Received: July 13, 2016 Revised: September 22, 2016

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Langmuir Finally, there exist limited studies on the fabrication of porous, monodisperse acrylamide-based microspheres because of their slow polymerization rate.32 Polyacrylamide (pAAm) is commonly used in multiple applications, including protein separation,33 water−oil separation,34 and drug release,35 because of its mechanical strength,36 hydrophilic nature, and strong antifouling properties37 and thus represents a desirable choice of polymer for biomacromolecular conjugation. However, pAAm-based microspheres in previous studies are either polydisperse24,25 or macroporous with the help of poly(ethylene glycol) (PEG), which introduced core−shelltype structures via phase separation.38 Therefore, pAAm-based microsphere systems represent a challenging space that can highlight the unique advantages of the batch-processing-based micromolding technique. Our approach to addressing these challenges centers on a simple and robust micromolding fabrication scheme based on surface tension-induced droplet formation in micropatterned poly(dimethylsiloxane) (PDMS) molds.39 Specifically, as shown in the schematic diagram of Figure 1a, aqueous prepolymer solutions consisting of acrylamide (AAm), acrylic acid (AA), and N,N′-methylenebis(acrylamide) (MBAM) are used to fill the wells of micropatterned PDMS molds. Next, hydrophobic wetting fluid consisting of hexadecane and a photoinitiator (2-hydroxy-2-methylpropiophenone) is pipetted onto the molds to allow surface tension-induced droplet formation of the prepolymer solution.40,41 Here the wetting fluid preferentially wets the PDMS surfaces of the microwells, causing a surface tension-induced pressure difference between the microwells’ edges and centers that pushes prepolymer solution toward the centers to form spherical droplets.40 Although in principle different microwell shapes can be used to form spherical droplets, the high surface area of edges of crossshaped microwells make this shape desirable for rapid and consistent spherical droplet formation.40 The molds are then irradiated with low-intensity (8 W) 365 nm UV light, which splits photoinitiator molecules into reactive radical species and initiates radical chain polymerization. To illustrate the dropletformation process, we present the bright-field micrographs of Figure 1b. Initially empty PDMS molds (left) are filled with prepolymer solution (middle). Upon the addition of wetting fluid, spherical droplets are spontaneously formed via surface tension minimization (right). As shown in the bright-field micrograph and accompanying distribution plot of Figure 1c, the resulting polymerized microspheres are highly uniform in dimensions as evidenced by the low coefficient of variation of 2.5% (Figure 1d). In this report, we first demonstrate the robust nature of the simple micromolding technique in the controlled fabrication of uniform p(AAm-co-AA) microspheres over a range of parameters. We also demonstrate the chemical functionality of the carboxylates in the as-prepared p(AAm-co-AA) microspheres by coupling them with the primary amines of a fluorescent marker via EDC/NHS chemistry. We then utilize this fluorescent marker as a semiquantitative measure to optimize the efficiency of the EDC/NHS reaction over a wide range of concentrations and ratios for hydrogel microparticle platforms. Finally, we show that the p(AAm-co-AA) microspheres possess porous network structures leading to improved protein conjugation kinetics via the EDC/NHS scheme using bright red fluorescent protein R-phycoerythrin (R-PE). Combined, these results demonstrate facile fabrication− conjugation schemes for uniform p(AAm-co-AA) hydrogel

Figure 1. Micromolding-based fabrication of p(AAm-co-AA) microspheres. (a) Schematic diagram of the fabrication procedure. (b) Bright-field micrographs of the PDMS mold at various stages of microsphere fabrication. Scale bars represent 500 μm. (c) Micrograph of the as-prepared microspheres and distribution plot of microsphere diameters. The scale bar represents 200 μm.

microspheres with chemical functionality and macroporous 3D network structures for improved biomacromolecular conjugation. We envision that the results and methodologies presented in this study could be readily extended to manufacture a wide range of functional hydrogels for biosensing and other applications.

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EXPERIMENTAL SECTION

Materials. Acrylamide (AAm) (99.9%), hydrogen chloride (HCl), 1-ethyl-3-(3-(dimethylamino)propyl)carbodiimide HCl (EDC), Nhydroxysuccinimide (NHS), 2-(4-morpholino)ethanesulfonic acid (MES), hexadecane, 2-propanol, sodium phosphate monobasic anhydrous (99%), sodium phosphate dibasic anhydrous (≥99%), Tween 20, poly(dimethylsiloxane) (PDMS) elastomer kits (Sylgard 184), and sodium hydroxide (NaOH) were purchased from Thermo Fisher Scientific (Waltham, MA). Acrylic acid (AA) anhydrous (180− 200 ppm MEHQ inhibitor, 99%), 2-hydroxy-2-methylpropiophenone (Darocur 1173, photoinitiator), saline sodium citrate (SSC) buffer (20× concentrate, molecular biology grade), and glucosamine HCl were purchased from Sigma-Aldrich (St. Louis, MO). N,N′B

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Langmuir Methylenebis(acrylamide) (MBAM) was purchased from EMD Millipore (Billerica, MA). Fluorescein glycine amide (FGA) was purchased from Setareh Biotech (Eugene, OR). R-Phycoerythrin (RPE) was purchased from Anaspec Incorporated (Fremont, CA). Poly(ethylene glycol) (PEG, MW 8 kDa) was purchased from Research Organics Incorporated (Cleveland, OH). All chemicals were analytical grade and used without further purification. Fabrication of p(AAm-co-AA) Microspheres via Micromolding. The pAAm and p(AAm-co-AA) microspheres in this study were fabricated according to methods in recent reports41 with minor modifications. Briefly, the compositions of the aqueous prepolymer solutions were 0−27.5% (w/v) AAm, 0−19% (w/v) AA, 0.1−2.5% (w/v) MBAM, and 0−2% (w/v) PEG porogen (MW 8 kDa), and the composition of the wetting fluid was 99% (v/v) hexadecane and 1% (v/v) Darocur 1173 photoinitiator. As shown in Figure 1a, prepolymer solution was placed into a micropatterned PDMS mold (225 wells per mold), which was formed with Sylgard 184 containing 10% (w/w) curing agent following overnight incubation at 65 °C on a silicon master template. Bubbles in the microwells were removed by rubbing the surface of the mold with a disposable pipet tip. Excess prepolymer solution was removed via pipetting, and the wetting fluid was then placed on top of the mold in order to lead to surface tension-induced droplet formation. To prevent rapid evaporation of the prepolymer solution, this process was carried out in a humidity chamber with at least 90% relative humidity.20,42 The mold was then carefully placed on an aluminum mirror (Thorlabs, Newton, NJ) and exposed to lowintensity 365 nm UV light with an 8 W hand-held UV lamp (Spectronics Corp., Westbury, NY) for 15 min to initiate radical chain polymerization. The microspheres were collected via pipetting, transferred to a microcentrifuge tube, and rinsed to remove any wetting fluid and unreacted chemicals as follows: mixing the microspheres in 2-propanol by pipetting, allowing them to settle to the bottom, and removing the supernatant. After rinsing at least three times with 2-propanol, the rinsing procedure was repeated at least three times with deionized water containing 0.05% (v/v) Tween 20 and at least twice with 20 mM MES buffer (adjusted to pH 6 with 1 N NaOH) containing 0.05% (v/v) Tween 20 or deionized water. The pH of each buffer solution was measured with a standard pH probe (Thermo Scientific Orion 5-Star Plus Benchtop Meter, Thermo Scientific, Beverly, MA) prior to use. EDC/NHS Activation of the Microspheres. To convert the carboxylate groups in the microspheres into reactive NHS ester groups, 0−400 mM EDC and 0−400 mM NHS were added to microspheres in 20 mM MES buffer (pH 6) containing 0.05% (v/v) Tween 20, mixed via pipetting, and placed on a rotator for 15 min at room temperature. Unreacted EDC and NHS were removed by rinsing the microspheres with 20 mM MES buffer (pH 6) containing 0.05% (v/v) Tween 20 at least three times and with 2× SSC (adjusted to pH 6, 7, 8, or 9 with 1 N HCl or 1 N NaOH), 20 mM sodium phosphate buffer (adjusted to pH 6, 7, 8, or 9 by varying the ratio of monobasic and dibasic sodium phosphate and adjusting with 1 N NaOH), or 20 mM phosphate-buffered saline (pH 7.2) at least twice, using the rinsing procedure mentioned above. Fluorescent Labeling of Microspheres with FGA. For FGA conjugation, pAAm and p(AAm-co-AA) microspheres activated with EDC/NHS (roughly 50 particles) were reacted with 100 μM FGA for 5−120 min on a rotator at room temperature in the buffers mentioned in the previous section (EDC/NHS activation of the microspheres). The reaction was then quenched with 0.1 M excess glucosamine hydrogen chloride for 15 min on a rotator. Unreacted FGA was removed by rinsing the microspheres three times with a 50% (v/v) 2propanol 50% (v/v) deionized water mixture and twice with 5× SSC buffer (pH 7) containing 0.05% (v/v) Tween 20 using the rinsing procedure described above. R-PE Protein Conjugation with the Microspheres. For R-PE conjugation, pAAm and p(AAm-co-AA) microspheres (roughly 50 particles) activated with EDC/NHS were reacted with 1−4 μM R-PE for up to 24 h on a rotator at room temperature. To further aid in mixing, the microspheres were mixed by pipetting in 15−30 min intervals for the first 2−4 h of reaction as necessary. The reaction was

then quenched with 0.1 M excess glucosamine HCl for 15−30 min on a rotator. Unreacted R-PEs were removed by rinsing the microspheres at least seven times with 5× SSC buffer (pH 7) with 0.05% (v/v) Tween 20 using the rinsing procedure described above. Image Analysis. Fluorescently labeled and R-PE-conjugated microspheres were visualized with an epifluorescence microscope (Olympus BX51 equipped with a DP70 microscope digital camera, Center Valley, PA) and a confocal microscope (Leica DMIRE2 equipped with a TCS S9 scanner, Wetzlar, Germany) in 5× SSC buffer (pH 7) with 0.05% (v/v) Tween 20. Epifluorescence micrographs were obtained with a 10× objective lens using a standard green (UN31001) or red (U-N31002) filter set (Chroma Technology Corp., Rockingham, VT). Confocal micrographs were obtained with a 10× objective lens at 488 and 543 nm excitation for the R-PE-conjugated microspheres. Diameters and total fluorescence intensities of the microspheres were examined using ImageJ image analysis software.43 Total fluorescence intensities were calculated by multiplying the average fluorescence intensity of a microsphere by its associated area from epifluorescence micrographs. Error bars represent the standard deviation from at least five randomly selected spheres per condition examined.

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RESULTS AND DISCUSSION Robust Micromolding-Based Fabrication of p(AAmco-AA) Microspheres with Tunable Features. First, as shown in Figure 2, we demonstrate that the micromolding process is a robust route to fabricating uniform p(AAm-co-AA) microspheres over a large range of fabrication parameters. For this, we filled cross-shaped micromold patterns with prepolymer solutions of varying compositions and concentrations of AAm, AA, and MBAM, covered the filled micromolds with a hydrophobic wetting fluid containing photoinitiator to induce the formation of spherical droplets, and polymerized the microspheres with UV light. We then collected and washed the cross-linked microspheres and examined them via bright-field microscopy. First, the micrographs in Figure 2a show that p(AAm-co-AA) microspheres with highly uniform dimensions can be fabricated over a large range of total monomer contents, from 5 to 30% T. The AAm:AA:MBAM ratio was kept constant at 15:4:1 in this set of results. As shown in the left-most image of Figure 2a, microspheres fabricated with 5% T are quite transparent (arrows and dotted circle), as expected from the low polymer content and minimal difference in the refractive index (1.349 at 23 °C for 8% pAAm gel)44 from that of water (1.332 at 23 °C).44 These spheres are also larger than the spheres prepared with 10 and 30% T (224 μm diameter vs 185 and 188 μm, respectively), presumably because of the higher degree of swelling from the lower cross-linking density.45,46 In addition, Elliot et al. have shown that significant intramolecular cyclization of AA monomers can occur during the polymerization of cross-linked polymer systems with highly dilute prepolymer solutions.47 This pendant cyclization should also contribute to the increased swelling by decreasing the amount of cross-linking in the mesh.47 Next, Figure 2b shows that microspheres fabricated with various monomer to cross-linker ratios at fixed 2% AA and 12.5% T possess uniform dimensions for each condition yet display different colors and sizes. In the leftmost image of Figure 2b, spheres prepared with a high cross-linker content (4:1) are dark in color, likely because of the formation of macropores.32,48 Specifically, bundling of the polymer chains at high cross-linker contents has been reported to increase the mesh size.32,48 Next, as shown in the middle and rightmost micrographs of Figure 2b, microspheres with decreasing crossC

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brittle nature of poly(acrylic acid) (pAA)50 and the need for another component such as AAm to provide ductility.36 Meanwhile, volumetric swelling ratios of up to 20 indicate that pAAm and p(AAm-co-AA) microspheres can hold up to 95% water content in the swollen state (section 1, Supporting Information). Note that microspheres fabricated with a particular prepolymer composition may not retain that composition upon polymerization. For example, Riahinezhad et al. have shown that the monomer reactivity ratios, defined as the rate constant of homopolymerization over the rate constant of cross-propagation, of AAm and AA are 1.33 and 0.23, respectively, in AAm-AA copolymer systems.49 This indicates that the next monomer to be incorporated into a growing polymer chain is more likely to be an AAm monomer than an AA monomer, and thus not all of the AA in a prepolymer solution may be incorporated within the microspheres upon polymerization or polymer chains may form blocks of repeating AA when AAm is fully consumed (composition drift).51 The robustness of the micromolding process we examined in Figure 2 stems from the inherently batch-processing-based nature of this technique, which offers several unique advantages over microfluidic processes. For instance, micromolding can accommodate the use of monomer systems with slow polymerization rates, without the need for surfactants, delicate flow control, or viscosity tuning.52,53 Our simple micromolding procedure can also accommodate precise particle size control from less than 20 μm to greater than 250 μm simply by altering the volume of the microwells and the wettability of the wetting fluid and prepolymer solutions on the PDMS surfaces.19 Smaller particles with diameters below 1 μm can also be fabricated with micromolding by introducing simple techniques such as solvent evaporation.19 In addition, the uniformity of the particles fabricated with this process is evidenced by the consistently small standard deviations of the particle diameters for all of the conditions examined. In summary, the results shown in Figure 2 illustrate the robust nature of our micromolding process in fabricating uniform p(AAm-co-AA) microspheres. Chemical Functionality of p(AAm-co-AA) Microspheres. We next show that AA in the prepolymer solution is uniformly incorporated into the microspheres, providing abundant carboxylate functionality (Figure 3). For this, we utilized EDC/NHS chemistry to covalently couple primary amine groups from the small fluorescent molecule fluorescein glycine amide (FGA, MW 404.38 Da) to the microspheres’ carboxylates, as shown in the schematic diagram of Figure 3a. A more detailed description of the EDC/NHS reaction is presented in Scheme S1 in the Supporting Information. Specifically, microspheres prepared with varying AA contents were first activated with excess EDC/NHS to convert the carboxylates to NHS esters. Upon washing to remove unreacted EDC/NHS, the microspheres were exposed to 100 μM FGA to form stable amide linkages. We then washed away unreacted FGA from the microspheres and examined their fluorescence via epifluorescence microscopy. First, the bright-field and epifluorescence micrographs in Figure 3b show p(AAm-co-AA) microspheres fabricated with fixed AAm and MBAM and varying AA contents upon reaction with 5 mM EDC and 5 mM NHS and then with 100 μM FGA for 15 min each. The first two negative control conditions in Figure 3b show that microspheres lacking either AA or EDC/ NHS activation do not fluoresce significantly. In contrast,

Figure 2. p(AAm-co-AA) microspheres prepared with various fabrication parameters. (a) Bright-field micrographs of microspheres fabricated with varying 5−30% T but a constant AAm:AA:MBAM ratio of 15:4:1. (b) Bright-field micrographs of microspheres fabricated with 12.5% T (fixed 2% AA and varying AAm and MBAM) in the prepolymer solution. (c) Bright-field micrographs of microspheres fabricated with 20% T in the prepolymer solution with varying AAm and AA (fixed 1% MBAM). (d) Microspheres fabricated with 12.5% T in the prepolymer solution, swollen in deionized water (top row), and then dried via vacuum desiccator (bottom row). Scale bars represent 200 μm.

linker content down to 99:1 are also fabricated consistently but are larger and more transparent (arrow and dotted circle), likely because of the decrease in cross-linking density and the corresponding increase in swelling. Next, the micrographs in Figure 2c show that microspheres prepared with varying AA contents can also be fabricated consistently. Here, microspheres were fabricated with fixed 1% MBAM, 20% T, and with varying AA and AAm content from 0 to 19 and 19−0% of the 20% T respectively. Increasing the amount of AA in the prepolymer solution led to an increase in the diameter of the microspheres, likely because of the less efficient polymerization of AA.49 Moreover, microspheres prepared with higher AA content are darker in appearance, which could again be the result of macropore formation.32,48 As AA content reaches 10−19% of the 20% T, the microspheres become more nonuniform and begin to crack, confirming the D

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more hydrolysis-resistant intermediate than does EDC alone.54 Importantly, these results indicate that the fluorescence of the microspheres is due to the covalent coupling of FGA molecules with the microspheres’ carboxylate groups with minimal nonspecific adsorption. Next, the bottom two rows of Figure 3b show that microspheres fabricated with increasing AA content display increasing fluorescence upon EDC/NHS activation and FGA labeling, up to 10% T (i.e. 48.8% (v/v) of polymerizable monomer components in the prepolymer solution). This trend is attributed to the increased incorporation of AA in the microspheres with increasing AA in the prepolymer solution, allowing for more carboxylate groups within the microspheres to react with EDC/NHS and subsequently with FGA. In addition, the diameters of the microspheres fabricated with 0− 10% AA increased from 161 to 195 μm, respectively. This increase can be attributed to the poor incorporation efficiency of AA mentioned earlier49 (Figure 2) and the relative increase in the monomer to cross-linker ratio for microspheres fabricated with increasing AA content, leading to decreased cross-linking density and increased swelling. Finally, to gain a better understanding of the chemical reactivity of the carboxylate groups, we further examined the fluorescence intensity of the microspheres shown in Figure 3b via image analysis (ImageJ, Experimental Section). Figure 3c shows that the microspheres’ total fluorescence intensity increases in a nonlinear fashion with increasing AA content in the prepolymer solution, even after accounting for the microspheres’ slight size increase with increasing AA in the prepolymer solution. This nonlinear trend can be attributed to a few factors. First, AA may not be incorporated as readily into the microspheres as is AAm, and thus linearly increasing the AA content in the prepolymer solution may not correspond to the same linear increase in the carboxylate content in the polymerized microspheres.49 Alternatively, most or all of the AA in the prepolymer solution may be incorporated into the microspheres, yet the difference in reactivity ratios between AAm and AA may lead to AAm monomers being consumed more quickly than are AA monomers. This could lead to long chains of repeating AA after AAm is fully consumed. As a result, the carboxylates in chains composed of repeating AA monomers could be subject to steric hindrance for the EDC/ NHS activation and/or FGA labeling. Taken together, the results in Figure 3 indicate that p(AAm-co-AA) microspheres contain uniformly incorporated carboxylate functional groups (from AA) that covalently couple with primary amines via EDC/NHS chemistry. pH-Dependent Kinetics of FGA Labeling with p(AAmco-AA) Microspheres via EDC/NHS Chemistry. As shown in Figure 4, we next examined the kinetics of the FGA labeling reaction with EDC/NHS-activated p(AAm-co-AA) microspheres under various pH conditions. The pKa of FGA’s primary amine is ∼9.6 (cf. lysine amine pKa ≈ 10.25, Nterminal amine pKa ≈ 8−9),55,56 making FGA an adequate model small molecule for studying protein conjugation via EDC/NHS chemistry. For this, we first reacted p(AAm-co-AA) microspheres with 5 mM EDC/NHS and removed unreacted EDC/NHS from solution upon reaction. We then labeled the EDC/NHS-activated microspheres with 100 μM FGA under various pH conditions for varying time points and quenched the reaction with excess glucosamine before washing the microspheres thoroughly and analyzing their total fluorescence.

Figure 3. Fluorescent labeling of p(AAm-co-AA) microspheres with FGA. (a) Schematic diagram showing fluorescent labeling via EDC/ NHS chemistry. (b) Bright-field and epifluorescence micrographs of fluorescently labeled microspheres with fixed 10% AAm, 0.5% MBAM, and varying AA content in the prepolymer solution. Scale bars represent 200 μm. (c) Total fluorescence intensity vs AA content in the prepolymer solution.

microspheres containing AA that are activated with EDC/NHS and labeled with FGA display highly uniform fluorescence (upper rightmost column, Figure 3b), indicating that the AA groups are uniformly incorporated into the microspheres and are chemically functional, and that small fluorescent marker FGA (MW 404.38 Da) can readily diffuse through the microspheres within 15 min. Meanwhile, microspheres activated with EDC show only lower fluorescence, whereas those activated with NHS show only minimal fluorescence (Figure S1, Supporting Information). This result confirms that EDC is necessary in activating carboxylate groups whereas NHS enhances the amine coupling reaction by providing a E

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Figure 4. Kinetics of the FGA labeling reaction with EDC/NHS-activated p(AAm-co-AA) microspheres under various pH conditions. (a) Total fluorescence intensity vs time for microspheres labeled with FGA via EDC/NHS chemistry at pH 6−9 and accompanying micrographs at 120 min. (b) Epifluorescence micrographs of microspheres reacted at pH 8 at each time point.

First, the total fluorescence plot in Figure 4a shows that the reactions at pH 8 and 9 proceeded rapidly, reaching 75 and 85% of each condition’s maximum fluorescence (2 h), respectively, within the first 15 min. In contrast, the reactions at pH 6 and 7 proceeded slowly, reaching only 33 and 53%, respectively, within 15 min (dashed red vertical line). This contrast in the apparent conjugation rate is likely attributed to the decrease in the primary amine’s reactivity with decreasing pH from the progressive loss of nucleophilic character necessary to attack the carbonyl carbon of the NHS ester groups in the microspheres. At pH 6 and 7, the primary amines of FGA are mostly protonated (i.e., positively charged) and are not reactive for the nucleophilic acyl substitution reaction with NHS esters, whereas at pH 8 and 9 the primary amine retains sufficient electron density to react rapidly. In addition, the fluorescence intensities of the pH 8 and 9 conditions reach a plateau by 2 h, whereas the fluorescence intensities of the pH 6 and 7 conditions continue to increase slowly. The fluorescence upon the 2 h reaction also varied for each pH, with pH 9 yielding the highest fluorescence and pH 6, 7, and 8 yielding 40, 67, and 90% of the final fluorescence intensity at pH 9, respectively, as further illustrated in the fluorescence micrographs on the right of Figure 4a. Here, the competing pHdependent hydrolysis of NHS esters during reaction likely contributes to the rapid plateauing of the pH 8 and 9 conditions, which should reduce the number of NHS ester groups available to react with FGA. These results are consistent with previous reports describing the increase in the NHS ester hydrolysis rate with increasing pH.23,57,58 Note that for each condition the fluorescence intensities both within and among microspheres are uniform, as partially evidenced by the consistently small error bars, illustrating the consistency of our fabrication−conjugation scheme. Finally, the epifluorescence micrographs in Figure 4b taken at various time points show uniform fluorescence over the particle dimensions without higher fluorescence around sphere edges,

particularly at the 5 min time point, illustrating the rapid mass transfer of small FGA molecules within the microspheres. Combined, these results indicate that FGA is a suitable model small molecule for examining the conjugation reaction via the EDC/NHS scheme with p(AAm-co-AA) microspheres without diffusion limitation. EDC/NHS Activation Conditions. Next, we examined optimal EDC/NHS concentrations and ratios for the fluorescent labeling of FGA with p(AAm-co-AA) microspheres, as shown in Figure 5. For this, we activated p(AAm-co-AA) microspheres with various concentrations and ratios of EDC/ NHS and subsequently labeled them with 100 μM FGA upon washing.

Figure 5. Total fluorescence intensity plot of p(AAm-co-AA) microspheres labeled with FGA with various concentrations and ratios of EDC/NHS. F

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Langmuir First, the open columns in the semilog total fluorescence intensity plot in Figure 5 show that p(AAm-co-AA) microspheres activated with 5× theoretical molar excess EDC/NHS over AA in the prepolymer solution (Section 4 in Supporting Information) show increasing fluorescence with increasing EDC, up to 0.4 mM EDC and 0.1 mM NHS. Microspheres activated with EDC/NHS in the 5 and 100 mM ranges displayed a similar increasing trend in fluorescence intensity with increasing EDC concentration. In fact, in each concentration range, doubling the EDC/NHS ratio from 1:1 to 2:1 to 4:1 led to a similarly proportional increase in fluorescence intensity. This increase is likely due to the greater number of EDC molecules allowing for a faster apparent reaction rate. Although the reaction should continue beyond 15 min, the time frame chosen should yield sufficient NHS ester groups for the subsequent fluorescent labeling while minimizing the hydrolysis of the NHS esters and O-acylisoureas. Thus, for a given concentration of NHS, it makes sense for the microspheres activated with the highest EDC concentration (400 mM EDC, 100 mM NHS) to yield the highest fluorescence intensity. Meanwhile, microspheres activated with 100 mM EDC and either 200 mM NHS or 400 mM NHS yielded substantially low total fluorescence intensities (Figure S3) of 1.9 × 107 and 1.2 × 107 AU, respectively (data not shown). This decrease in fluorescence intensity with increasing NHS concentration may be a result of the adsorption of NHS groups near the carboxylates (via hydrogen bonding or electrostatic interaction), thus interfering with the formation of O-acylisourea.59 When comparing the various concentration ranges, microspheres activated with 400 mM EDC and 100 mM NHS displayed only 5× higher fluorescence intensity than did microspheres activated with 20× less EDC and NHS (20 mM EDC and 5 mM NHS). Moreover, microspheres activated with 20 mM EDC and 5 mM NHS displayed the same fluorescence intensities as did microspheres activated with 100 mM EDC and 100 mM NHS. These results indicate that a 4:1 ratio of EDC/NHS yields the highest amine coupling efficiency and that increasing the EDC/NHS concentration to above the 5 mM range yields diminishing returns. As such, the results in Figure 5 are in agreement with Sam et al.’s results on EDC/ NHS activation on acid-modified silicon surfaces, which found that the EDC/NHS activation of carboxylates in the 5 mM range yielded the most NHS ester groups with a minimal formation of less reactive anhydride or inactive N-acylurea side products.60 However, our results are somewhat contrary to Wang et al.’s results on the EDC/NHS activation of pAA polymer brushes on silicon surfaces, which found that the activation of pAA brushes with 0.1 M EDC and 0.2 M NHS was optimal.61 Finally, the consistently small error bars (Figure 5) and uniform fluorescence within and among microspheres (Figure S3) for all of the conditions examined again demonstrate the robustness of our fabrication−conjugation scheme. In summary, these EDC/NHS concentration studies indicate that carboxylate groups activated with high EDC/NHS concentration ranges at 4:1 or a higher EDC/NHS ratio lead to the most amine coupling on microscale hydrogel platforms. Conjugation of Large R-PE Proteins with p(AAm-coAA) Microspheres via EDC/NHS Chemistry with Minimal Nonspecific Adsorption. As shown in Figure 6, we next examined protein conjugation with the p(AAm-co-AA) microspheres via EDC/NHS chemistry using a bright red fluorescent protein R-phycoerythrin (R-PE) as a model protein. R-PE’s

Figure 6. R-PE conjugation with p(AAm-co-AA) microspheres via EDC/NHS chemistry. (a) Schematic diagram of R-PE conjugation. (b) Epifluorescence micrographs of R-PE-conjugated microspheres. The top row shows microspheres with varying AA activated with (+/+) or without (−/−) EDC/NHS and then conjugated with R-PE at pH 8. The bottom row shows EDC/NHS-activated microspheres conjugated with R-PE at pH 7, 8, or 9. Scale bars represent 200 μm. (c) Total fluorescence intensity plot for R-PE-conjugated microspheres.

large size (MW 240 kDa, hydrodynamic diameter (Dh) ≈ 11 nm) and multiple primary amine sites on the outer surface41,62,63 (Figure S4) make the protein an ideal model for studying biomacromolecular conjugation and the polymer mesh size.41,64 For this, we activated p(AAm-co-AA) and pAAm microspheres with excess EDC/NHS for 15 min and then with R-PE overnight under various pH conditions, as shown in the schematic diagram of Figure 6a. We then utilized epifluorescence microscopy to examine the R-PE-conjugated microspheres. First, as shown for the three negative control conditions of Figure 6b, microspheres lacking either AA or EDC/NHS activation displayed minimal fluorescence upon reaction with R-PE, indicating the selectivity of the EDC/NHS scheme and the nonfouling nature of pAAm and p(AAm-co-AA) microspheres. Microspheres prepared with high AA content (top rightmost image) also showed minimal fluorescence, indicating minimal nonspecific binding by electrostatic attraction with carboxylates under the pH conditions used. Specifically, G

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Figure 7. R-PE conjugation kinetics and confocal micrographs of p(AAm-co-AA) microspheres. (a) Fluorescence intensity vs time for R-PE conjugation (2 μM) with 12.5% T microspheres fabricated with 0−2% PEG porogen (MW 8 kDa) in the prepolymer solution and accompanying confocal micrographs in the centers of the microspheres. (b) Fluorescence intensity vs time for R-PE conjugation (1−4 μM) with 12.5% T microspheres fabricated with 0% PEG in the prepolymer solution and accompanying confocal micrographs in the centers of the microspheres. Scale bars represent 200 μm.

These results are in good agreement with the FGA labeling results in Figures 3 and 4. The uniformity of the fluorescence in the R-PE-conjugated microspheres matches that of the FGAlabeled microspheres, suggesting that the microspheres are macroporous and contain uniformly distributed carboxylate groups. Combined, these results confirm that EDC/NHS chemistry with p(AAm-co-AA) microspheres is a simple, straightforward, and selective route to protein conjugation with minimal nonspecific protein adsorption. Porous Network Structures of p(AAm-co-AA) Microspheres via R-PE Conjugation Kinetics. Finally, we carried out a thorough R-PE conjugation kinetics study to examine the mesh size of the p(AAm-co-AA) microspheres, as shown in Figure 7. For this, we fabricated p(AAm-co-AA) microspheres with varying concentrations of an inert poly(ethylene glycol) porogen (PEG, MW 8 kDa) that showed the efficient formation of macropores at low content in other pAAmbased systems.64,66−68 Upon activation with EDC/NHS and thorough washing, we conjugated the microspheres with varying concentrations of R-PE for up to 8 h at pH 8, quenching the reaction with excess glucosamine at each time point. We then imaged the microspheres via epifluorescence (Figure S5) and confocal microscopy to examine the kinetics and spatial fluorescence profiles, respectively. First, as shown in the total fluorescence intensity plot of Figure 7a, EDC/NHS-activated 12.5% T p(AAm-co-AA) microspheres fabricated with 0−2% PEG porogen in the prepolymer solution all exhibited similar R-PE conjugation kinetics, quickly reaching 80% of the final concentration within 2 h (dotted red line) and starting to plateau by 4 h. The similar

although both carboxylates and R-PE are net negatively charged under the pH conditions used (carboxylate pKa ≈ 4.3, R-PE isoelectric point ≈ 4.5−5),65 positively charged patches on the R-PE surface may still be attracted to carboxylates on the microspheres. The results shown here indicate that this type of electrostatic attraction is negligible. In contrast, the results in the bottom row of Figure 6b show that the microspheres activated with EDC/NHS and reacted with R-PE at pH 7, 8, or 9 display bright and uniform fluorescence among the microspheres, demonstrating the facile nature of protein conjugation with our microspheres using EDC/NHS chemistry. Furthermore, each microsphere displays uniform fluorescence throughout its dimensions, providing preliminary evidence that the R-PEs have diffused fully throughout each microsphere. Here, the pH 8 and 9 conditions displayed similar fluorescence intensity in contrast to the lower fluorescence of the pH 7 condition. These differences in fluorescence intensity among the conditions enlisted are quantified via image analysis in the fluorescence intensity plot of Figure 6c. Figure 6c confirms that the negative control conditions display minimal fluorescence whereas the pH 7, 8, and 9 conditions display substantially higher fluorescence. However, the pH 8 and 9 conditions displayed 33% higher fluorescence than did the pH 7 condition, likely because of less protonation of the R-PE’s N-terminal amines (pKa ≈ 7.8) and lysines (pKa ≈ 10.5) at higher pH, leading to more nucleophilic acyl substitution of the NHS esters (Figure 6a).56 Furthermore, the pH 8 and 9 conditions displayed similar levels of fluorescence, suggesting that the R-PEs had sufficiently deprotonated primary amines to react with the NHS esters under both pH conditions. H

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Langmuir R-PE kinetics and final fluorescence for the three PEG porogen conditions suggest that R-PE molecules can quickly diffuse through the polymer networks and reach the NHS ester groups within the microspheres for each of the conditions examined. In other words, this result implies that p(AAm-co-AA) microspheres are sufficiently macroporous to facilitate the diffusion of R-PE without porogen. This is likely due to the low polymer content and low cross-linking density arising from the inefficient incorporation of AA monomers during polymerization mentioned earlier (Figure 2). The accompanying confocal micrographs of the centers of the microspheres in Figure 7a illustrate the rapid diffusion of R-PE into the microspheres’ centers within 30 min for each condition and saturation in the microspheres’ centers by 2 h, thus providing concrete evidence of the microspheres’ macroporous nature with or without PEG porogen. In our coupled diffusion−reaction system of R-PE conjugation with the p(AAm-co-AA) microspheres, negatively charged carboxylates are converted to reactive NHS esters, anhydrides, (a smaller number of) unstable O-acylisoureas, and inactive N-acylureas within their polymer networks60 upon activation with EDC/NHS (Scheme S1). Driven by the concentration gradient, net negatively charged R-PE proteins (MW ≈ 240 kDa, Dh ≈ 11 nm, pI ≈ 4.5−5.1)41,62,65 containing multiple primary amines (from N-termini and lysine groups) will first react with the O-acylisourea or NHS ester groups present on the microspheres’ surfaces to form amide linkages after diffusing through the boundary layer. These large proteins will continue to react with NHS esters or O-acylisourea groups as they diffuse through the polymer networks toward the microsphere centers. Concurrent with this reaction is the hydrolysis of the O-acylisourea and NHS esters into carboxylates. Furthermore, the pores within the polymer networks will become narrower as more R-PE’s react and further restrict R-PE diffusion via steric hindrance. In Figure 4, we have already shown that the amine coupling reaction between NHS ester groups and FGA molecules occurs on the order of minutes to hours, whereas the time scale of R-PE diffusion into polymeric microspheres depends heavily on the polymer mesh size.68 As such, we expect that initially R-PEs should react quickly with the reactive groups in the microspheres, resulting in a high initial apparent rate of reaction reflected in a steep increase in the total fluorescence intensity with time. As the reaction continues, R-PEs should experience more hindered diffusion as bound R-PEs occupy space within the pores. This slower diffusion in addition to the hydrolysis of NHS esters should result in the slower observed apparent rate of reaction.41,69,70 Next, in Figure 7b, we conjugated 1, 2, or 4 μM R-PE with p(AAm-co-AA) microspheres without PEG porogen to examine the effect of concentration on the resulting kinetics. As shown in the fluorescence intensity plot of Figure 7b, the 4 μM R-PE condition (open squares) exhibited the highest final total fluorescence, whereas the 1 μM R-PE (open circles) and 2 μM R-PE (filled triangles) conditions displayed 40 and 60% of the 4 μM R-PE condition’s final total fluorescence, respectively. Because there is theoretically at least a 10-fold molar excess of NHS ester groups over R-PEs at 1 μM concentration (section 4 in the Supporting Information), an increase in the final total fluorescence intensity with increasing R-PE concentration should be expected. This large excess should lead to apparent pseudo-first-order kinetics with respect to R-PE, with an increasing concentration of R-PE leading to a faster apparent

reaction rate. This is reflected in the high initial rates of each condition examined in Figure 7b, with the 2 and 4 μM conditions displaying 1.7 and 2.5 times the initial rate of the 1 μM condition, respectively (on the basis of the initial fluorescence intensities). However, the rapid rate of decay for each condition indicates a significant deviation from pseudofirst-order kinetics. This observation can be attributed to the diffusion limitation of R-PEs, the hydrolysis of NHS esters mentioned earlier, and/or multiple reactive sites (i.e., primary amines) on the R-PEs. As more R-PE’s are conjugated with the microspheres, their pores will become narrower and NHS ester groups will continue to hydrolyze, making subsequent conjugation more difficult. It is important to note here that although the half-life of NHS esters is ∼1 h at pH 8 and room temperature,58 the high theoretical excess of NHS ester groups in the microspheres compared to R-PEs in solution should allow reaction to continue well past 1 h, albeit at reduced rates. In addition, as mentioned earlier, R-PEs have multiple primary amines (i.e., N-termini as well as lysine’s amines) throughout their surfaces, which may lead to the consumption of multiple NHS ester groups per single R-PE molecule and hence an increase in the consumption of NHS ester groups during conjugation. Furthermore, the large size of R-PE will likely sterically hinder neighboring NHS ester groups from reacting with amine groups from other R-PEs. These contributing factors may help to explain the deviation from pseudo-firstorder kinetics in the kinetic profiles of the three conditions examined. Meanwhile, the accompanying confocal micrographs of the centers of the microspheres in Figure 7b further confirm the rapid diffusion of R-PEs in the microspheres within 30 min. However, the 1 μM condition shows microspheres with dimmer centers at 30 min and 1 h compared to those under the 2 and 4 μM conditions, corresponding to their fluorescence profiles in the fluorescence vs time plot. This result is in accordance with the lower driving force for the diffusion of RPE into the microspheres at lower concentration. Finally, note that the 2 μM condition in Figure 7b (dark triangles), which independently mirrors the 0% PEG condition in Figure 7a (dark triangles, separate batch of experiments), yields similar fluorescence intensities with the corresponding condition in Figure 7a at each time point examined (differing by 13% or less), indicating the robustness of our fabrication−conjugation scheme. These R-PE kinetics results highlight the utility of our fabrication−conjugation scheme utilizing p(AAm-co-AA) microspheres and EDC/NHS chemistry over those used in our recent studies. Briefly, microspheres were fabricated using the same micromolding technique with 10% poly(ethylene glycol) diacrylate (PEGDA, MW 700 Da) and 0.5% chitosan (CS) in the prepolymer solution and were reacted with 2 μM R-PEs via rapid tetrazine-trans-cyclooctene (Tz-TCO) chemistry.41 In this more diffusion-limited system, the microspheres reached 50% reaction completion within 2 h and 80% completion by 8 h. Compared to these microspheres, the p(AAm-co-AA) microspheres in the present work displayed ∼80% maximum fluorescence within 2 h for the same concentration of R-PE used, thus demonstrating the more macroporous nature of p(AAm-co-AA) microspheres. In another study, CS-PEGDA microparticles were surfacefunctionalized with cysteine-modified tobacco mosaic viruses (TMVs) and similarly reacted with 2 μM R-PEs via Tz-TCO chemistry.70 Although this system is also diffusion-limited, the R-PEs diffuse only through the boundary layer and several I

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confirm the reaction conditions of carboxylate functional groups in hydrogel microsphere formats, which otherwise vary widely in the literature.27−29 Next, R-PE conjugation studies showed that the carboxylates of p(AAm-co-AA) microspheres can readily couple with primary amines on large proteins with minimal nonspecific adsorption. Finally, rapid diffusion and reaction of R-PE within 30 min and 2 h, respectively, into p(AAm-co-AA) microspheres established the highly macroporous nature of p(AAm-co-AA) microspheres. In addition, the apparent conjugation reaction rate of large unmodified proteins with p(AAm-co-AA) microspheres described in this work is substantially higher than those in our recent works.41,68,70 In conclusion, these findings highlight the potential of p(AAm-co-AA) microspheres for biomacromolecular conjugation via EDC/NHS chemistry for various biosensing applications.

micrometers of TMV-assembled regions, with minimal diffusion through polymeric networks. Here, the fluorescence rapidly reached its maximum value within 3 h. Compared to these results, our p(AAm-co-AA) microspheres reached their maximum fluorescence with the full penetration of R-PE within 2 h for the same concentration of R-PE. This further highlights the macroporous nature of our microspheres. Finally, our group has also recently estimated the diffusion coefficient of R-PEs and the mesh size of micromolding-based CS-pAAm microsphere systems.68 Here, CS-pAAm microspheres fabricated with 2.5% PEG porogen (MW 8 kDa) in the prepolymer solution reached full penetration of R-PEs upon 24 h conjugation via rapid Tz-TCO reaction, and the estimated mesh size of such microspheres was roughly 39 nm.68 Compared to the CS-pAAm microspheres, the p(AAm-coAA) microspheres here reached full penetration of R-PE within 2 h, suggesting that the mesh size of p(AAm-co-AA) microspheres is substantially larger than 39 nm. These results further demonstrate the utility of our p(AAmco-AA) microspheres in biomacromolecular conjugation over our previous platforms and other methods. The robust micromolding method allows for the microspheres’ highly uniform dimensions, the uniform incorporation and distribution of carboxylate groups, and the macroporous nature of the polymer networks without the need for any porogens. These properties in turn allow for the complete diffusion and reaction (within 2 h) of large biomacromolecules (e.g., antibodies (MW ∼150 kDa) and R-PE (MW ∼240 kDa)) via EDC/NHS chemistry in a straightforward manner, without the need for additional modification or activation steps for the proteins to be conjugated.41 In contrast, polymer brush systems suffer from environment-dependent deformed brush configuration and collapse, requiring the careful tuning of reaction and working conditions.71 These results confirm that p(AAm-co-AA) microspheres are macroporous with full penetration of R-PE within 30 min and can be utilized for the conjugation of large biomacromolecules via EDC/NHS chemistry within 2 h.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.langmuir.6b02591. Volumetric swelling ratio calculations, EDC/NHS reaction mechanisms, bright-field and epifluorescence micrographs of FGA-labeled microspheres and negative controls, calculations of theoretical prepolymer contents, crystal structure model of R-PE, and epifluorescence micrographs of R-PE-conjugated microspheres (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Telephone: (617) 627-2195. Fax: (617) 627-3991. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We extend our special thanks to Dr. Chang Hyung Choi at Harvard University and Professor Chang-Soo Lee at Chungnam National University for providing silicon mastermolds and helpful advice on the micromolding-based fabrication technique. We gratefully acknowledge partial financial support from the Shirley & Stanley Charm Scholarship Fund (E.Y.L.) and the Tufts Faculty Research Fund.

CONCLUSIONS In this article, we demonstrated the facile fabrication of macroporous and carboxylate functional p(AAm-co-AA) hydrogel microsphere platforms for rapid biomacromolecular conjugation via EDC/NHS chemistry. First, we demonstrated the robust nature of the micromolding method in fabricating macroporous p(AAm-co-AA) microspheres with uniform dimensions and adjustable carboxylate functionality by simply varying the composition and concentration of the prepolymer solution. These fabrication results represent a significant innovation in the ability to fabricate monodisperse and highly macroporous polyacrylamide-based microspheres with abundant carboxylate chemical functionality by exploiting the unique features of our batch-processing-based micromolding technique. Next, we enlisted FGA labeling via EDC/NHS chemistry to show that p(AAm-co-AA) microspheres have abundant and uniformly distributed carboxylate functional groups. We further examined the utility of FGA as a small model molecule to study protein conjugation in the absence of diffusion limitation by examining the kinetics of the amine coupling reaction between carboxylates in p(AAm-co-AA) microspheres and primary amines. We then semiquantitatively determined the optimal EDC/NHS ratio to be 4:1 at high millimolar concentrations for the EDC/NHS activation of carboxylates and the subsequent amine coupling reaction. These EDC/NHS results clarify and

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DOI: 10.1021/acs.langmuir.6b02591 Langmuir XXXX, XXX, XXX−XXX

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DOI: 10.1021/acs.langmuir.6b02591 Langmuir XXXX, XXX, XXX−XXX