PEGMA-Based Microgels: A Thermoresponsive Support for Enzyme

Article pubs.acs.org/Macromolecules

PEGMA-Based Microgels: A Thermoresponsive Support for Enzyme Reactions Sepehr Mastour Tehrani,†,‡ Yijie Lu,‡ and Mitchell A. Winnik*,†,‡ †

Department of Chemical Engineering and Applied Chemistry, University of Toronto, 200 College Street, Toronto, ON M5S 3E5, Canada ‡ Department of Chemistry, University of Toronto, 80 St. George Street, Toronto, ON M5S 3H6, Canada S Supporting Information *

ABSTRACT: Thermoresponsive colloidal hydrogels (aqueous microgels) made from poly[oligo(ethylene glycol) methacrylate] (PEGMA) are an interesting class of biomaterials due to their sharp thermal transition and excellent biocompatibility. However, the inherent protein repellency of PEGMA has made the biofunctionalization of these microgels difficult and prevented them from reaching their full potential in applications such as a protein carrier. Here, we report the synthesis of thermoresponsive PEGMA microgels and the covalent attachment of horseradish peroxidase (HRP, as a model protein) to these microgels. We prepared our microgels by the precipitation copolymerization of 70 mol % OEGMA300 and 30 mol % methacrylic acid in water. The resulting microgels showed a volume phase transition temperature (VPTT) of ca. 60 °C in acidic buffers and in DI water but in neutral and basic buffers did not show a thermal response up to 75 °C. The direct immobilization of HRP to the activated carboxylic groups in the microgel backbone was unsuccessful, but using a diamine spacer with four ethylene glycol units, we were able to covalently attach this enzyme to our PEGMA microgels through bis-aryl hydrazone chemistry. HRP has its maximum activity at ca. 50 °C. At higher temperatures, the activity was reduced, but the microgel-bound enzyme showed less reduction in activity than the native enzyme and no change in activity associated with the VPTT. In addition, PEGMA microgels stabilized the attached enzymes against thermal denaturation. For example, our results showed that the enzyme immobilized on the PEGMA microgel lost its activity 3.4 times slower than the free enzyme in the first 5 h of annealing at 50 °C. The bioconjugation strategy introduced here could serve as a model for the covalent attachment of other biomacromolecules to the protein-repellent PEGMA microgels.

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INTRODUCTION Thermoresponsive colloidal hydrogels (aqueous microgels) exhibiting a volume phase transition temperature (VPTT) in aqueous media are often referred to as smart materials.1,2 Poly(N-isopropylacrylamide) (PNIPAM) is the most studied and most often used thermoresponsive polymer in the fabrication of aqueous microgels. PNIPAM microgels are fully swollen in water at room temperature, but a small increase in temperature causes them to shrink and show a transition at ca. 32 °C, which corresponds to the lower critical solution temperature (LCST) of the homopolymer solution in water. This unique property made PNIPAM microgels a desirable class of biomaterials with applications in enzyme immobilization,3 bioreactors,4 hyperthermia-induced drug delivery,5 cell carriers,6 and tissue engineering.7 However, despite its widespread application in biotechnology, PNIPAM has major drawbacks. These include hysteresis in its phase transition, and a significant influence of functional groups on its thermal behavior.8,9 There are also comments in the literature about the toxicity of PNIPAM microgels,8 but this may be due to residual monomer. © XXXX American Chemical Society

In a search for a suitable replacement for PNIPAM, Lutz and co-workers in a series of publications10−12 introduced polymers made from oligo(ethylene glycol) methacrylate (OEGMA) of different ethylene glycol (EG) chain lengths. These polymers, denoted PEGMA, show a range of transition temperatures, from 22 to 95 °C, depending on their EG length. Increasing the length of the pendant EG groups renders the polymer more hydrophilic, and its VPTT moves to a higher temperature. The copolymerization of OEGMA monomers of different EG lengths allows one to prepare microgels with a VPTT intermediate between those of the two pure components. For example, by the copolymerization of di(ethylene glycol) methacrylate (DEGMA, 95 mol %) and OEGMA475 (Mn ∼ 475 g mol−1, 5 mol %), microgels with a VPTT of 32 °C were prepared, resembling that of PNIPAM.9,13 By changing the feed ratio of these monomers, the VPTT of the resulting microgels could be adjusted to any temperature between those of Received: June 14, 2016 Revised: October 13, 2016

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DOI: 10.1021/acs.macromol.6b01270 Macromolecules XXXX, XXX, XXX−XXX

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Macromolecules

replacement for PNIPAM microgels as supports for enzymes, without reporting any relevant studies. Here, we report the synthesis of temperature- and pH-responsive PEGMA microgels and the conjugation of horseradish peroxidase (HRP) enzyme as a model protein to these microgels. These PEGMA microgels were synthesized by precipitation copolymerization oligo(ethylene glycol) methyl ether methacrylate (Mn ∼ 300 g/mol) (OEG4MA) and methacrylic acid (MAA) with poly(ethylene glycol) dimethacrylate (Mn ∼ 550) as a cross-linker. In a second step, we coupled tetra(ethylene glycol)ethyldiamine (TEGDA) to the MAA groups as a spacer of similar length to OEGMA monomer used (∼4 EG units). This spacer provided amino functionality on the PEGMA microgels that we used for the conjugation to HRP enzyme through bis-aryl hydrazone (BAH) chemistry. We examined the microgel-bound enzyme and compared its activity to the native enzyme at different pH values and temperatures. We also compare the reactivity of HRP bound to PEGMA microgels to that of HRP bound to poly(hydroxyethyl acrylamide) (PHEAA) over a similar range of temperatures, reported previously.27

PDEGMA and PEGMA475. The thermal transition of PEGMAbased microgels is very sharp and less susceptible to hysteresis than PNIPAM microgels. Apart from its useful thermal behavior, PEGMA shows excellent biocompatibility properties.12 It is well-known that linear PEG is water-soluble, nontoxic, biocompatible, and protein repellent. It is by far the most commonly used synthetic polymer in biotechnology.14 Several recent studies also verified the biocompatibility of PEGMA as “PEG analogues”.11,15 For examples, PEGMA copolymers incubated with the human hepatocellular carcinoma (HepG2) cells lines did not induce cell death, and cell viability was comparable with cells incubated with commercial linear PEG.16 In a mouse model, injected PEGMA-coated superparamagnetic iron oxide nanoparticles (SPION) exhibited long circulation times and excellent antibiofouling behavior.17,18 Grafting PEGMA brushes on different materials conferred excellent protein resistance to their surfaces. Surfaces such as gold, polystyrene, and silica coated with PEGMA brushes showed remarkable biocompatibility.19 Despite these benefits, targeted biofunctionalization of PEGMA microgels and dense PEGMA brushes remains a challenging task. The protein resistance of PEGMA hinders the attachment of target proteins to the activated functional groups incorporated into PEGMA copolymers. To improve the application of PEGMA-based structures in biotechnology such as microarrays, enzyme carriers, or biosensors, a rather sophisticated chemical design is required to attain a functionalizable polymer host and simultaneously keep the matrix, protein repellent.20 Finding a simple approach to decorate PEGMA chains with biomacromolecules is a subject of interest in recent years. Most studies on this topic have relied on the modification of the terminal −OH groups of hydroxylcapped PEGMA pendant chains. For example, the authors of refs 21 and 22 activated this terminal hydroxyl with disuccinimidyl carbonate and then attached biotins and used streptavidin as a capture protein to immobilize it on the biotin at the end of each OEGMA unit. In a different approach, Theato and colleagues23 synthesized PEGMA block copolymers through RAFT polymerization with a thyroxin at the initiating end and used the copolymer end group to attach biotin, which in turn was used to bind to streptavidin. Chen et al.24 reported that they were able to make dendritic structures by the ATRP copolymerization of OEGMA and N-acryloyloxysuccinimide (NAS), in which each methacrylate chain was end-capped with a bioreactive NHS moiety. Another elegant approach was reported by Hucknall et al.25 in which they removed water molecules in the vicinity of PEGMA chains through “vacuum desiccation” process to reduce the polymer hydration degree and enhance protein adsorption into the PEGMA brushes. Most of these techniques for the bioconjugation of PEGMAcoated surfaces are relatively complicated and not suitable for PEGMA microgels. Immobilization of proteins on PEGMA microgels introduces an extra level of complexity to the chemical design. In the literature, there are many reports on the bioconjugation of proteins to other microgels such as PNIPAM, whereas the covalent immobilization of protein on PEGMA microgels remains unexplored. While there are a few reports on the biofunctionalization of polymer particles such as poly(methyl methacrylate) sterically stabilized via PEG chains, there are as yet no examples of the direct bioconjugation to aqueous PEGMA microgels. In several review papers,19,26 PEGMA microgels were mentioned as a “future and potential”

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METHODS

Instrumentation. Electron Microscopy (EM). Transmission EM (TEM) images were obtained using a Hitachi H-7000 transmission electron microscope operating at 75 kV. Alternatively, TEM images were obtained from the transmission mode of a Hitachi S-5200 highresolution scanning electron microscope (SEM) running at 30 kV. SEM images were obtained at 1 kV. Microgel samples were washed with dilute aqueous HCl solution and redispersed in DI water and deposited on PELCO carbon-coated copper grids. Dynamic Laser Scattering (DLS). DLS measurements were performed on a Malvern Zetasizer Nano ZS instrument at a backscattering angle of 173°. For measurements at elevated temperatures, samples were allowed to equilibrate for 5 min after reaching the desired temperature. UV−Vis Spectroscopy. UV−vis spectra were recorded in either 0.2 or 1 cm path-length disposable cells using an Agilent Cary 300 spectrophotometer equipped with a 6 × 6 Peltier temperature controller and two internal temperature probes. Turbidity measurements were recorded 5 min after the microgel solutions (0.5 mg/mL) reached the desired temperature. Materials. Monomer oligoethylene glycol methyl ether methacrylate (Mn ∼ 300 g/mol) (OEGMA300), methacrylic acid (MAA), initiator ammonium persulfate (APS, 98%), cross-linker poly(ethylene glycol dimethacrylate) (Mn ∼ 550 g/mol) (PEGDMA550), diamine spacers ethylenediamine (EDA), hexamethylenediamine (HMDA), PEG-diamine (Mn ∼ 2000) (PEGDA2000), peroxidase from horseradish (HRP, type VI-A), 2,2′-azinobis(3-ethylbenzothiazoline-6sulfonic acid) diammonium salt (ABTS assay, 98% HPLC), hydrogen peroxide (30 wt % in H2O), N-(3-(dimethylamino)propyl)-N′ethylcarbodiimide hydrochloride (EDC, 98%), N-hydroxysuccinimide (NHS, 98%), 4-(4,6-dimethoxy-1,3,5-triazin-2-yl)-4-methylmorpholinium chloride (DMTMM, 96%), 4-formylbenzoic acid (97%), 6-chloropyridine-3-carboxylic acid (99%), hydrazine hydrate, anhydrous DMF (99.8%), potassium phosphate (monobasic and dibasic) (99%), hydrochloric acid and sodium hydroxide (1 N, HPLC standard), and trimethylamine (99%) were purchased from SigmaAldrich. Spacer tetra(ethylene glycol)ethyldiamine (Mn = 236.3 g/ mol) (TEGDA) was purchased from ChemPep Inc. Fluorescent dyes, NHS-rhodamine, and NHS-fluorescein (5/6-carboxyfluorescein succinimidyl ester, mixed isomers) were purchased from ThermoFisher Scientific. Succinimidyl 4-formylbenzoate (S-4FB) and succinimidyl 6hydrazinonicotinate acetone hydrazone (S-HyNic) were synthesized as described by Grotzky et al.28 B


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Macromolecules Table 1. Chemical Structures and Amounts of Diamines Used To Attach to PEGMA-MAA Microgels

fluorescein to label microgels with HMDA and PEGDA spacers. For the fluorescein labeling, we used NHS-fluorescein (1.9 mg, 4 μmol) dissolved in DMSO (50 μL). An aliquot of PEGMA-NH2 stock solution (100 μL, 1.0 mg microgel) in phosphate buffer (100 mM, pH 8.0) was diluted in phosphate buffer (350 μL, 100 mM, pH 8.0). The microgel and NHS-dye solutions were mixed together with a vortex mixer inside an amber tube. The reaction was continued for 4 h at room temperature, and then the mixture was washed extensively through repeated centrifugation and redispersion in PBS buffer (pH 7.4) until the supernatant became colorless. At the end, the PEGMArhodamine microgels were dispersed in PBS buffer (1 mL, pH 7.4). PHEAA-NH2 Microgel Coupling to HyNic. PEGMA-NH2 microgels (3 mg) were suspended in phosphate buffer (400 μL, 100 mM, pH 8.0). To this solution, an aliquot of HyNic (1.01 mg, 3.5 μmol) in DMSO (100 μL) was added. The reaction continued for 2 h at room temperature, and then unreacted HyNic was washed off the microgel mixture by repeated centrifugation and redispersion and in phosphate buffer (50 mM, pH 5.2). The microgels were redispersed in this buffer and diluted to a volume of 220 μL. An aliquot of this solution (20 μL) was withdrawn to quantify the number of HyNic per microgel, and the remaining 200 μL was reacted with the S-4FB-modified HRP (see the following). To evaluate the number of HyNic per microgel, 4formylbenzoic acid (80 μL, 50 mM) was added to 20 μL of the PHEAA-HyNic solution, and the evolution of the absorbance at 354 nm was followed by UV−vis (ε354 nm = 29 000 M−1 cm−1).28 HRP Coupling to S-4FB. HRP enzyme (0.9 mg, ca. 20 nmol) was dissolved in phosphate buffer (400 μL, 100 mM, pH 8.0). To this solution, an aliquot of S-4FB (30 μg, 120 nmol) in DMSO (10 μL) was added. The (S-4FB/HRP = 6) ratio was selected to avoid each HRP molecule having more than one 4FB group. The reaction continued for 2 h at room temperature, and then unreacted S-4FB was separated from the HRP by repeated spin filtering using a 10 000 MWCO spin filter and phosphate buffer (50 mM, pH 5.2). After extensive washing cycles, 4FB-modified HRP was suspended in phosphate buffer (220 μL, 50 mM, pH 5.2). An aliquot of this solution (20 μL) was withdrawn to quantify the number of 4FB per HRP molecule, and the remaining 200 μL was reacted with the HyNicdecorated PHEAA microgel. To evaluate the number of 4FB groups per HRP molecule, a solution of 2-hydrazinopyridine (80 μL, 500 μM) in phosphate buffer pH 5.0 was added to the HRP-4FB solution (20 μL), and the evolution of the absorbance at 350 nm was followed by UV−vis (ε350 nm = 24 500 M−1 cm−1).29 PEGMA-HyNic and HRP-4FB Conjugation. The PEGMA-HyNic solution (200 μL) and the HRP-4FB solution (200 μL) (described earlier) in phosphate buffer (pH 5.2) and an aqueous solution of aniline (40 μL, 100 mM) were mixed together in a UV cell. The evolution of absorbance at room temperature was monitored at 354 nm (ε354 nm = 29 000 M−1 cm−1) until the value reached an apparent plateau (1 h). At the end of the reaction, nonconjugated HRP-4FB was separated from conjugated PEGMA-HRP by centrifugation and redispersed in phosphate buffer, pH 6.8. This sample is denoted PEGMA-HRP and was stored at 4 °C. Determination of Enzymatic Activity. The Michaelis−Menten constant Km and the maximum reaction velocity Vm for the free and immobilized HRP were determined by monitoring the increase in absorbance in the ABTS assay at varying concentrations of the ABTS

Poly(oligoethylene glycol methyl ether methacrylate) (PEGMAMAA) Microgel Synthesis. Precipitation polymerization of OEGMA300 was conducted in a 250 mL three-necked round-bottom flask, equipped with a condenser, steel stirring rod with a Teflon paddle, and a nitrogen/chemical inlet. OEGMA300 (1 g, 3.3 mmol), MAA (86 mg, 1 mmol), PGDMA550 cross-linker (55 mg, 0.1 mmol), and sodium bicarbonate (10 mg, 0.11 mmol) were dissolved in Milli-Q water (95 mL) and transferred to the flask. The mixture was heated to 85 °C and bubbled with nitrogen for 45 min, and then APS initiator (16 mg, 70 μmol) dissolved in Milli-Q water (5 mL) was injected to the solution to start the polymerization. The reaction was allowed to proceed for 18 h under continuous nitrogen bubbling and 200 rpm mixing. At the end of the reaction, a homogeneous white microgel solution obtained was transferred to 100 kDa MWCO spin filters (Amicon, 15 mL). The microgel solution was washed extensively using DI water to remove all side products. After the final washing cycle, the microgel solution was redispersed in DI water to a final concentration of 9 mg/mL to serve as a stock solution. Attempted Bioconjugation to PEGMA-MAA Microgels Using EDC/NHS. PEGMA-MAA microgels (4 mg) were suspended in MES buffer (500 μL, 100 mM, pH 5.5). To this solution, a mixture of EDC/ NHS (50 mg/50 mg) in MES buffer (500 μL, 100 mM, pH 5.5) was added, and they mixed at room temperature for 20 min. Then this solution was quickly washed through two cycles of centrifugation and redispersion with PBS buffer (1X, pH 7.4). After the second centrifugation step and supernatant withdrawal, HRP enzyme (1 mg) in phosphate buffer (400 μL, 100 mM, pH 8.0) was mixed with the activated microgels and reaction continued for 3 h. The reaction mixture was purified extensively through repeated centrifugation and redispersion with phosphate buffer (50 mM, pH 6.8) until no enzymatic activity was observed from the supernatant. Attempted Bioconjugation to PEGMA-MAA Microgels Using DMTMM. PEGMA-MAA microgels (4 mg, 4 μmol COO−) were suspended in phosphate buffer (400 μL, 50 mM, pH 7.4) and mixed with a moderate excess of DMTMM (1.4 mg, 5 mmol) for 5 min. Then, HRP enzyme (1 mg) in phosphate buffer (500 μL, 200 mM, pH 8.0) mixed with the activated microgel solution for 4 h. The reaction mixture was purified extensively through repeated centrifugation and redispersion with phosphate buffer (50 mM, pH 6.8) until no enzymatic activity was observed from the supernatant. Diamine Spacer Attachment to PEGMA-MAA Microgels. For each experiment, PEGMA-MAA microgels (4 mg, 4 μmol COO−) were suspended in phosphate buffer (400 μL, 50 mM, pH 7.4) and mixed with a moderate molar excess of DMTMM (1.5 mg, 5.5 μmol) for 5 min. Then, a large molar excess of diamine spacer (EDA or HMDA or PEGDA2000 or TEGDA), as shown in Table 1, in phosphate buffer (1 mL, 200 mM, pH 7.8) was mixed with the activated microgels for 4 h (the pH of diamine solutions was adjusted with HCl prior to addition to the activated microgels). The reaction mixture was purified extensively through repeated centrifugation and redispersion with phosphate buffer (50 mM, pH 5.2). We refer to the coupling product with TEGDA as PEGMA-NH2 microgels. Fluorescent Dye Labeling. To label amine-containing microgels with a fluorescent dye, NHS-rhodamine (2.1 mg, 4 μmol) was dissolved in DMSO (50 μL). At some point during our experiments, we ran out of NHS-rhodamine in our laboratory and used NHSC


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Macromolecules substrate. To perform this assay, an aliquot of HRP solution (50 μL, 50 nM) in phosphate buffer (50 mM, pH 6.8), H2O2 (50 μL, 0.3 wt %, 88 mM), and varying amounts of ABTS solution (0−900 μL, 1.0 mM) in phosphate buffer (50 mM, pH 5.2) were mixed in a cuvette and diluted with phosphate buffer (50 mM, pH 5.2) to a total reaction volume of 1.00 mL. The kinetics of this assay was monitored by recording the change in the absorbance of the solution at 405 nm over the first 60 s of reaction. All the kinetic measurements were performed in a thermostated cell at 25.0 °C. The extent of product formation was monitored by measuring the increase in absorbance at 405 nm using ε405 nm = 36 800 M−1 cm−1. Km and Vmax values were calculated from the expression

ϑ=

Vmax[S] K m + [S]

85 °C well above the LCST of PEGMA300 and continued for 18 h using poly(ethylene glycol) dimethacrylate (PEGDMA, M ∼ 550, 2.3 mol %) as cross-linker. This recipe resulted in the formation of homogeneous microgels with a narrow size distribution as shown in the TEM image in Figure 1A. This

(1)

where [S] is the ABTS substrate concentration and ϑ is the initial rate. Microsoft Excel was used for curve fitting. Activity Measurement at Different pH. The activity of free and immobilized HRP was studied in phosphate buffers over a pH range of 4−9 using pyrogallol as the substrate. To perform this assay, an aqueous solution of pyrogallol (500 μL, 3 wt %), phosphate buffer (400 μL, 100 mM), an HRP solution (free or immobilized) (50 μL, 50 nM) in phosphate buffer (50 mM, pH 6.8), and H2O2 (50 μL, 0.3 wt %, 88 mM) were mixed in a UV cell. The enzyme kinetics was studied by recording the increase in the absorbance of the solution at 420 nm over the first 120 s of reaction. Activity Measurements at Different Temperatures. In this experiment, the rate of change in absorbance of each sample was recorded in a thermostated Peltier cell. ABTS solution (500 μL, 1.0 mM) in phosphate buffer (pH 5.2, 50 mM) and phosphate buffer (400 μL, pH 5.2, 50 mM) were mixed together inside the UV cuvette and placed inside the cell, which was equipped with an internal probe for precise temperature control. After the desired temperature was reached at each step, 50 μL of (50 nM) native or immobilized HRP in phosphate buffer (pH 6.8, 50 mM) plus H2O2 solution (50 μL, 0.3 wt %, 88 mM) was added to the mixture, and the rate of change in absorbance (over the first 1 min) was measured at 405 nm. Enzyme Thermal Stability. For the thermal stability experiment, free and immobilized enzyme solutions at a concentration of (50 nM) were annealed at 50 °C in phosphate buffer (pH 6.8, 50 mM) in the absence of substrate. Sample withdrawal was performed every 30 min. These aliquots (50 μL) were mixed with the kinetic test solution [ABTS solution (500 μL, 1.0 mM) in phosphate buffer (pH 5.2, 50 mM), phosphate buffer (400 μL, pH 5.2, 50 mM), H2O2 (50 μL, 0.3 wt %, 88 mM)], and the rate of change in absorbance at 405 nm (over 1 min) was measured at 25 °C. The decay rates of the enzymatic residual activity were fitted with an exponential decay model using the first point obtained at 50 °C (5 min annealing) as t0 = 5 min.

(%) residual activity = α exp(−kdecayt )

Figure 1. (A) TEM image of PEGMA-MAA microgels obtained from precipitation polymerization at 85 °C. (B) LCFM image of PEGMANH2 microgels decorated with a rhodamine dye and dispersed in PBS buffer (pH 7.4).

(2)

where α is a prefactor.

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TEM image (see also a lower magnification TEM image in Figure S1A, Supporting Information) shows that the microgels have a narrow size distribution. DLS measurements of these PEGMA-MAA microgels in DI water gave a hydrodynamic diameter dz = 694 nm and a PDI of 0.046 (Figure S1B). pH Response of PEGMA-MAA Microgels. To quantify the content of carboxylic acid in the PEGMA-MAA microgel, we used simultaneous conductometric and potentiometric titration, in which the microgels were first treated with a small excess of NaOH (to pH 11) and then back-titrated with 0.1 N HCl solution (see Figure S2A). From both titrations, we found 31 mol % (1 μmol/mg microgel) of carboxylic acid functionality. From the plot of ionization degree vs pH (Figure S2B), we infer that with increasing pH, the initial deprotonation occurs at ca. pH 4.6, and full ionization is reached only at pH 10. We use this information to calculate the

RESULTS AND DISCUSSION Microgel Preparation. In this research, we studied the synthesis and characterization of temperature- and pHresponsive PEGMA-MAA microgels and the bioconjugation of HRP enzyme to these microgels. The PEGMA-MAA microgels were prepared by aqueous precipitation copolymerization of oligo(ethylene glycol) methyl ether methacrylate (OEGMA300) and methacrylic acid (MAA). In precipitation polymerization, all ingredients including the monomers, initiator, and cross-linker are dissolved in water, and at the polymerization temperature (e.g., 40−90 °C), the propagating polymer chains become insoluble and start to precipitate. The precipitates serve as nuclei for further microgel growth. The reported LCST for the PEGMA300 homopolymer in water is around 61−68 °C.12,19 The polymerization was carried out at D


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Macromolecules distribution of pKa values of the −COOH groups in the microgel (Figure 2).30 This plot shows that there is a

Figure 2. Distribution of pKa values for the −COOH groups in the PEGMA-MAA microgel sample calculated from the titration curve shown in Figure S2A.

distribution of pKa values, ranging from ca. 6.2 to 8.4. The distribution arises from the high MAA content of the microgels such that the ionization of some of the −COOH groups generates −COO(−) ions that influence the pKa of acid groups in their proximity. The pH response of the PEGMA-MAA microgel sample in phosphate buffers of different pH was studied by DLS. These data (Figure 3A) show that with increasing pH the microgels swell and their hydrodynamic diameters increase. This increase levels off after pH 8, suggesting that further ionization of the −COOH groups in the microgel does not lead to an increase in swelling. The pH-responsive swelling behavior is related to the deprotonation of carboxylic acid groups, in which the influx of counterions causes an increase of osmotic pressure inside the microgel. This acts as a driving force for additional water to diffuse into the microgels.31 Temperature Response of PEGMA-MAA Microgels. The thermal behavior of linear PEGMA in water has been studied extensively. Lutz12 reported the transition temperatures of various PEGMA derivatives. In the case of PEGMA300, the reported transition temperature varies between 61 and 68 °C. However, factors such as polymer molecular weight, comonomer polarity, and measurement buffer (type of salt, concentration, and pH) influence the transition temperature. For microgels, the network cross-link density also impacts the detected VPTT. Most of the previous studies on the thermal response of PEGMA microgels were carried out on microgels prepared by copolymerization of DEGMA and OEGMA475, designed to mimic the thermal behavior of PNIPAM. Here, we report the thermal response of our PEGMA300MAA microgels in DI water, NaCl solution, and phosphate buffers of different pH. Microgel samples at a concentration of 0.5 mg (microgel)/mL (buffer) were examined by turbidity measurements at 380 nm. The results of these measurements are shown in Figure 4. With increasing temperature, microgels in DI water, in NaCl solution (150 mM), and in PB (50 mM) at pH 5.2 started to become turbid at temperatures higher than 40 °C. However, a more dramatic change in turbidity occurred at 60 °C, which we assign as the VPTT of our microgels. One can also observe in Figure 4 that degree of microgel deswelling

Figure 3. Effect of pH on (A) PEGMA-MAA microgel size (green plot) obtained from the cumulant analysis of DLS autocorrelation decay data and (B) relative enzymatic activity of native HRP (red plot) and PEGMA-HRP (blue plot) obtained from pyrogallol assay at 25 °C. The enzymatic activities were normalized to their maximum at pH 9.0.

Figure 4. Turbidity measurements as a function of temperature for PEGMA-MAA microgel soutions in various aqeuous media (c = 0.5 mg/mL). For each measurement, the microgel solutions were allowed to equilibrate for 5 min at each temperature, and then the corresponding absorbance was recorded at 380 nm.

and increase in turbidity in PB pH 5.2 is higher than in DI water. According to our DLS measurements, dz = 663 nm for the PEGMA-MAA microgels in PB at pH 5.2 at 25 °C, decreasing to dz = 232 nm at 65 °C (see Figure S3). In contrast, microgels in neutral and basic buffers such as PB (50 mM, pH 7.4), PBS buffer (50 mM, pH 7.4) (results not shown), and PB (50 mM, pH 8.0) did not show any thermal response up to 75 E


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Macromolecules °C. These observations can serve as a guide for the prediction of the thermal response of functional PEGMA-MAA microgels in different pH regions. Upon heating from 25 to 65 °C, the PEGMA microgels suspended in PB pH 8.0 did not show any significant change in their apparent diameter (dz = 847 nm), but there was a notable increase in their PDIs, from (PDI = 0.056) at 25 °C to (PDI = 0.3) at 65 °C. We presume that the ionization degree of poly(methacrylic acid) constituent of our PEGMA-MAA microgel was high enough in basic buffers to keep the microgels swollen at the higher temperatures. Another set of results on the influence of composition on the LCST of PEGMA and PEGMA-MAA copolymers was reported by Becer et al.32 They studied the thermal response at different molar ratios of PEGMA and MAA (0−100%) in water. They showed that the incorporation of more MAA into the PEGMA chains shifted the LCST of the polymer to lower temperatures. In their system, the polymer was dissolved in water and the carboxyl groups were largely protonated. They attributed their observations to hydrogen bonding between the −COOH groups and the EG repeat units of the polymer. These intramolecular hydrogen-bonding interactions reduced the polymer swelling in water. Direct Immobilization of Proteins to PEGMA-MAA Microgels. We studied two types of coupling reactions in an attempt to immobilize HRP (as a model protein) directly to our microgels. The −COOH groups in the PEGMA-MAA microgels (31 mol %) were first activated with either EDC/ NHS or DMTMM. Then, the enzyme was introduced into the reaction vessel to react with the activated microgels, presumably through its lysine amino groups. All of these attempts to attached HRP to the carboxylated PEGMA microgel in this way yielded such tiny amounts of conjugation product that the signal for protein attached to the microgel was barely distinguishable from background. More details about these experiments are presented in the Supporting Information. This observation accords very well with previous reports on the protein-repellent properties of PEGMA microgels or brushes.33 Surfaces with grafting densities as low as 0.2−0.3 PEGMA chains/nm2 were shown to be strongly protein repellent.18 In our PEGMA-MAA microgel, the activated −COOH groups in the microgel backbone are buried among the dense PEG pendant groups. It appears to be difficult for proteins to displace the water molecules that hydrate the PEG chains in order to reach the activated −COOH groups. Attachment of Spacers to the PEGMA-MAA Microgel. To render the functional groups more accessible for bioconjugation, we modified our PEGMA-MAA microgels with the different diamine spacers (Table 1). We used DMTMM chemistry to attach the spacer molecules to the activated −COOH groups of the microgels. The time for the activation reaction was kept short (5 min) so that not all the −COOH groups were activated. In this way we aimed to preserve a large fraction of the −COOH content of the microgel to maintain the pH-responsiveness of the final microgels. The diamine spacers were then added in a large molar excess in order to minimize any possible cross-linking. We also found a very specific set of reaction conditions that gave a maximum spacer coupling yield. It entailed carboxylate activation via DMTMM (1.4 mol equiv) in pH 7.4 phosphate buffer (50 mM) for 5 min, followed by the addition of diamine in pH 7.8 phosphate buffer (200 mM). In our hands, deviations from these conditions (e.g., different pH) resulted in lower coupling yields.

We used four types of diamine spacers (EDA, HMDA, PEGDA2000, and TEGDA) to functionalize our PEGMA microgels. After each reaction, the microgels were purified from excess reagents by repeated centrifugation−redispersion. Then the amine content of the microgel was assessed by a reaction with an excess amount of NHS-rhodamine dye. In this way, we estimated the efficiency of the spacer coupling reaction. When the PEGMA-MAA microgels were functionalized with EDA, we found about 20% conversion of the −COOH groups. These microgels preserved their size distribution and colloidal stability (Figure S4A). We were unable to immobilize HRP on these microgels using BAH chemistry. A similar coupling reaction with HMDA resulted in a somewhat lower coupling conversion. Only 16% of the −COOH groups were transformed to amine functionalities. Unfortunately, this spacer caused the microgels to lose their colloidal stability and aggregate. The aggregates can be seen in the laser confocal fluorescence microscopy (LCFM) image of these microgels in PBS buffer shown in Figure S4B. As a much longer spacer, we used a poly(ethylene glycol) diamine PEGDA2000. Here only 1% of the −COOH groups were derivatized with this spacer, and this spacer also caused the microgels to aggregate and lose their colloidal stability (Figure S4C). The best results were obtained with tetraethylene glycol diamine (TEGDA) as the spacer. The 4 EG units of TEGDA are similar in length to the 4−5 EG units of the OEGMA300 pendant groups of our PEGMA-MAA microgel. In a previous study of poly(hydroxyethyl acrylamide) microgels with an ethylenediamine spacer, we were able to use 1H NMR to quantify the amine content. For the PEGMA−TEGDA conjugate, we were unable to use NMR to quantify the extent of TEGDA conjugation because of peak overlap with the OEGMA peaks from the microgel. As a consequence, we used a different approach to quantify the amine content of the microgels. We reacted the amine groups of the spacer with rhodamine-NHS and measured the UV−vis absorbance of the dye-labeled microgel. This assay assumes quantitative reaction with rhodamine-NHS and the same molar extinction coefficient for the microgel-bound dye as for the free dye in solution. In this way we found that 8% of the COOH groups reacted with TEGDA. We refer to the PEGMA−TEGDA microgel sample as PEGMA-NH2. The rhodamine dye provided a fluorescence signal for confocal microscopy, and an LCFM image of this sample is presented in Figure 1B. The PEGMA-NH2 microgels appear to be colloidally stable with a diameter of ca. 700 nm. We measured the electrophoretic mobilities of the PEGMAMAA microgels before and after modification with the TEGDA spacer, in 100 mM phosphate buffer, both at pH 7.8 (DMTMM conjugation pH) and at pH 5.0. At pH 7.8 both microgels gave significant negative values (−2.0 × 10−8 m2/(V s), equivalent to an apparent zeta-potential of −15 mV). For the microgel with TEGDA spacer, the net negative charge is consistent with the conversion of only a small fraction of −COOH groups to amino groups. These results show that the modification of the microgels with diamine spacer did not change the surface charge of microgels. At pH 5, the electrophoretic mobilities were negative (−0.6 × 10−8 m2/(V s)) but very small in magnitude, suggesting that the microgel surfaces had no significant net charge. Enzyme Immobilization. We chose bis-aryl hydrazone (BAH) chemistry 34,35 to attach HRP to PEGMA-NH 2 microgels. To proceed, PEGMA-NH2 microgels were initially decorated with S-HyNic as shown in Scheme 1A. We used an F


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Scheme 1. (A) Chemical Modification of PEGMA-MAA Microgels with the TEGDA Spacer through DMTMM Coupling To Form PEGMA-NH2, Followed by the Reaction of This Amine-Containing Microgel with NHS-HyNic To Form PEGMA-HyNic; (B) Modification of HRP with NHS-4FB in PB (pH 8.0) To Form HRP-4FB; (C) Conjugation of HRP-4FB to PEGMA-HyNic in PB at pH 5.2 Using Aniline as a Catalysta

This conjugation step was followed by recording the absorbance of the solution at 354 nm (ε354 nm = 29 000 M−1 cm−1). After 1 h of conjugation, the enzyme loading was 1 nmol (44 μg) HRP/mg microgels. a

reaction (using aniline as the catalyst) as PEGMA-HRP. This is the material used in the experiments described below. Quantifying the Enzyme Loading. By monitoring the formation of BAH bonds through their absorbance at 354 nm (Figure S5A, ε354 nm = 29 000 M−1 cm−1), we could quantify the number of enzymes attached to the microgels. This analysis gave a value of 3 nmol of HRP attached to 3 mg of PEGMAHyNic microgel (1 nmol HRP/mg microgel, dry weight). We verified this amount with two separate measurements. First, we analyzed the difference between the concentration of free HRP molecules prior to (17.5 nmol) and after (14.5 nmol) incubation with the microgels. This analysis was based on a concentration calibration curve for HRP at 403 nm. The difference was also 3.0 ± 0.1 nmol of immobilized HRP (1.0 nmol HRP/mg microgel). Second, we examined the absorbance of the PEGMA-HRP microgel conjugates at 403 nm, correcting for the background absorbance of the microgel itself. This analysis gave a value of 2.9 ± 0.1 nmol of HRP, also 1.0 nmol HRP/mg microgel. In a previous publication,27 we described the use of similar chemistry to attach HRP to PHEAA microgels with dz = 800 nm. The enzyme loading reported for PHEAA, obtained by monitoring BAH bond formation at 354 nm, was 0.5 nmol (22 μg) HRP/mg microgel, half the HRP content of the PEGMAMAA microgels reported here. There are several possible reasons for this difference. Both samples contained similar amounts of HyNic per mg of microgels. For the enzyme

excess of S-HyNic (ca. 14 equiv) based on the amine content of the microgels and a 2 h reaction time to convert a fraction of the amine groups to HyNic groups (see Table S1). After extensive washing cycles of sedimentation redispersion to remove excess reactant, the HyNic-modified PEGMA microgels were titrated with 4-formylbenzoic acid. In this way, we found 2.8 nmol of HyNic groups per mg of microgel. In a separate tube, HRP was modified with S-4FB (Scheme 1B). Here the goal was to attach on average one 4FB group per enzyme, presumably to a lysine amino group. After purification of the modified enzyme, it was titrated with 2-hydrazinopyridine, as monitored by UV absorption. Here we found on average 1.18 4FB groups per HRP molecule. In the conjugation reaction PEGMA-HyNic microgels were treated with an excess of HRP-4FB enzymes, using aniline as a catalyst, at pH 5.2 (Scheme 1C). The reaction was monitored by the growth in UV absorbance at 354 nm (Figure S5A). The reaction was allowed to continue for 1 h at 25 °C, at which the increase in UV signal leveled off. In a separate experiment, we followed the bioconjugation kinetics without using aniline as a catalyst. These results are plotted in Figure S5B. Here the conjugation reaction took 14 h to reach a plateau in the UV signal. This is an important result to keep in mind for bioconjugation reactions of temperature-sensitive proteins which cannot survive long incubation times at room temperature. We denote the product obtained in the conjugation G


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Macromolecules conjugations reaction with the PEGMA-HyNic microgels, we used aniline as a catalyst for the BAH coupling reaction.29 The catalyst increased the reaction yield and reduced the reaction time by more than a factor of 10, as shown in Figure S5. We were unaware of aniline as a catalyst when we synthesized the PHEAA-HRP microgels. The two types of microgels also differ in the extent to which they are swollen for both the S-HyNic reaction at pH 8 and the enzyme coupling at pH 5. The PEGMA-MAA microgels, which contain an excess of carboxylate groups, are ionized and highly swollen at pH 8. Titration measurements showed that the PHEAA-NH2 microgels are largely unprotonated at pH 8.27 In contrast, the PEGMA-MAA microgels are not ionized to a significant degree at pH 5 (ca. 10%, Figure S2B), whereas the PHEAA-NH2 microgels are largely ionized at this pH. Ionization not only promotes swelling but changes the chemical environment inside the microgel and may affect the reactivity of the functional groups. HRP itself has an isoelectric pH (IP) of 7.2. Thus, it is positively charged at pH 5 and may have a lower propensity to diffuse into the PHEAA-NH2 microgels, which are also positively charged. The cross-link density of the microgels should also affect the extent of protein diffusion into the microgel. The PEGMA microgels were prepared with 2.5 mol % cross-linker in the synthesis, whereas the PHEAA microgels were prepared with 6 mol % cross-linker. From this perspective, the PEGMA microgels should have a lower crosslink density and a more open pore structures, allowing more HRP to diffuse inside the microgel. All of these factors likely play a role in determining the level of HRP loading of the microgels. The extents of conversion for the two types of microgels are collected in Table S1. HRP Kinetic Studies. The enzyme kinetics of PEGMAHRP was studied, and its catalytic activity was compared to that of the native enzyme. The measurements were performed at pH 5.2, at varying concentrations of the substrate ABTS [2,2′azinobis(3-ethylbenzothiazoline-6-sulfonic acid) diammonium salt] and at constant concentrations of H2O2 and HRP. The kinetic parameters, Km, Vmax, and kcat, were obtained from fitting the kinetic data to the Michaelis−Menten model (Figure S6). The results from experiments carried out at 25 °C are summarized in Table 2. From these data, we found a 3-fold decrease in the catalytic activity of immobilized HRP (kcat = 88 s−1) compared to the native HRP (kcat = 266 s−1). On the other hand, the similarity in their Km values shows that there is no mass transfer limitation for the substrates to access the microgel-bound enzymes compared to native enzyme. In our previous studies, we showed that upon the covalent immobilization of HRP on PHEAA microgels through BAH chemistry, the enzymatic activity of HRP remained unchanged. For the sake of comparison, the kinetic parameters of native and PHEAA-immobilized HRP from our previous study are also presented in Table 2. According to this table, the kcat of PHEAA-HRP (250 s−1) is very similar to that native HRP (266 s−1). Therefore, in comparison with the PEGMA microgel (88 s−1), PHEAA is a more efficient support for preserving the catalytic activity of immobilized HRP. While we are the first to report enzyme immobilization on PEGMA microgels, there are rather a limited number of examples in the literature on the attachment of enzymes to linear PEGMA polymers. Lutz and colleagues36 attached trypsin to a linear copolymer of DEGMA and PEGMA475 and showed this conjugate has a higher enzymatic activity than native trypsin.

Table 2. Kinetic Parameters of Native and Immobilized HRP from Fitting the ABTS Assay Data with Michaelis−Menten Modela sample native HRPb PEGMAHRPb native HRPc PHEAAHRPc

Km [μM]

Vmax [μM min−1]

turnover number kcat [s−1]

catalytic efficiency (kcat/Km) [s−1 mM−1]

254 ± 8

40.5 ± 1.0

266d

1047

272 ± 23

13.2 ± 0.4

88

324

276

31

258d

935

318

30

250

786

a

The assay was carried out in a constant concentration of H2O2 and HRP and varying concentration of ABTS dye in phosphate buffer (pH 5.2) at 25 °C. bMeasurements were carried out at 2.5 nM HRP. cData reported in ref 25, where measurements were carried out at 2.0 nM HRP. dThe difference (3%) in kcat values of native HRP measured here and that reported in ref 25 is considered to be a normal variation in measurements performed many months apart under slightly different experimental conditions.

Effect of pH on the Enzymatic Activity. We examined the effect of pH on the enzymatic activity of free and immobilized HRP. The experiments were carried out in phosphate buffer at different pH values ranging from 4 to 9. For these experiments we used pyrogallol as the substrate instead of ABTS. The results of this study are plotted in Figure 3B. The relative activities for the native and immobilized HRP are obtained from the normalization of activities with respect to their maximum at pH 9.0. Decreasing the pH reduces the enzymatic activities of both samples, and both samples showed a similar trend in the pH range 9 to 7. At lower pH, however, PEGMA-HRP showed a more drastic reduction in its activity compared to native HRP. This change correlates with the decrease in the microgel size with decreasing pH as seen in Figure 3A. A similar behavior has been observed for enzymes immobilized on different types of pH-responsive microgels (either cationic or anionic), in which the enzymatic activity of immobilized enzymes decreased in the collapsed state of the microgels.37,38 In pH-invariant supports for enzyme immobilization, such as chitosan beads,39 the relative activities of immobilized and free HRP were found to be similar over most pH ranges (4 to 9). In those studies, the polymer-supported enzyme appeared to be even more robust and active in alkaline solutions with a pH > 10. Effect of Temperature on the Enzymatic Activity. The measured VPTT of our PEGMA-MAA microgel in the enzyme assay buffer (pH 5.2) was ca. 60 °C. We studied the activity of native HRP and immobilized HRP (PEGMA-HRP) at different temperatures in the range of 10−75 °C in the assay buffer. The results of these experiments are plotted in Figure 5. At temperatures below 50 °C, the relative activity of native HRP parallels that of PEGMA-HRP. The similarity in the trends of enzyme activity indicate that the retarded activity of the PEGMA-supported HRP persists over this temperature range. In our experiment, native HRP showed its maximum activity at 50 °C, while the maximum activity of PEGMA-HRP shifted to a higher temperature (i.e., 55 °C), close to the onset of the VPTT of the microgel. An interesting result from our experiments is that above 55 °C the activity of the microgelbond HRP showed a significantly slower reduction rate compared to native HRP, well into the temperature range H


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Thermal Stability of Immobilized Enzyme. We studied the thermal stability of native and immobilized HRP by annealing their solutions at 50 °C. It took 5 min for the solutions originally at room temperature to reach 50 °C. The first aliquot was taken at this point, with subsequent aliquots taken at 30 min intervals. After cooling each aliquot to 25 °C, we measured their kinetics. The results of this study are presented in Figure 6. These activity data for both samples were

Figure 5. Enzymatic activity of native HRP (red diamonds) and PEGMA-HRP (blue squares) at different temperatures in PB (pH 5.2). The data were obtained from the ABTS assay by recording the increase in abosrbance at 405 nm. The error bars represent the range of miximum and minimum values for three replicate measurements. The two set of data were plotted on different scales to facilitate comparison of the temperature response of the two systems.

where one expects the microgel to have undergone its collapse transition and turn off the activity of immobilized enzyme. There is considerable variation in the literature about the influence of the LCST or VPPT on the activity of enzymes bound to thermoresponsive polymers. Whether the enzyme activity at and above the LCST is suppressed or enhanced appears to depend on the polymer, the enzyme, and the substrate examined. For example, Park and Hoffman showed that the enzymatic activity of β-galactosidase attached to PNIPAM hydrogel beads was “switched off” above the LCST of PNIPAM.40,41 These authors argued that above the polymer LCST the hydrogel beads became dehydrated and collapsed. As a result, mass transfer and access of the substrate to the enzyme were suppressed. In a more recent paper, Ballauff and colleagues42 reported the immobilization β-galactosidase on PS−PNIPAM core−shell microgels. They observed an almost 3-fold increase in the enzymatic activity of immobilized enzymes at temperatures higher than the LCST of PNIPAM. This unusual observation was attributed to a higher hydrophobic interaction above the polymer LCST, which enhanced β-galactosidase activity. Russell and co-workers43 attached chymotrypsin to the dual thermoresponsive copolymer poly(sulfobetaine methacrylamide-co-NIPAM) with the idea of “engineering” the enzyme properties. For example, by changing the solution temperature leading to polymer precipitation, the authors showed that the activity of enzyme decreased as a consequence of an increase in the Km parameter. This behavior was regarded as the result of increased steric hindrance around the enzyme and restricted access of substrates to the enzyme. In contrast, in the conjugate of a linear PEG-acrylate polymer and glucose oxidase (GOX), Liu et al.44 observed an increase in the activity of GOX above the polymer LCST. The authors hypothesized that upon the polymer chain shrinkage (to occupy a smaller volume) there would be more space (less steric hindrance) for the enzyme to access its substrate. In the current state of the art, there are as yet no universal governing principles to allow one to predict how immobilizing an enzyme on a thermoresponsive polymer support will affect enzyme activity or how the activity will change as the polymer undergoes a thermal response. There is a need for further experiments on this topic.

Figure 6. Residual activity of native HRP (red diamonds) and PEGMA-HRP (blue diamonds) after anealing the enzymes at 50 °C in PB (pH 6.8). The activity measurements were performed at 25 °C by recording the change in the absorbance of ABTS assay at 405 nm. Data were normalized to their activities 5 min after the annealing started, when the solution just reached 50 °C. The dashed lines are fits to an exponential decay profile.

normalized to their measured activity at t = 5 min. The results showed a significant difference between the residual activity of free and immobilized HRP. After 300 min annealing at 50 °C, PEGMA-HRP recovered 75% of its initial activity measured at t = 5 min, whereas native HRP recovered less than 40% of its initial activity. Both samples exhibited first-order decay, and the decay rates were fitted to an exponential profile (eq 2). At 50 °C, the rate constant for the decay of native HRP was found to be kdecay‑HRP = 3.4 × 10−3 min−1 and for immobilized HRP kdecay‑PEGMA = 1.0 × 10−3 min−1. These values show that the deactivation rate of native HRP at 50 °C is 3.4 times faster than that of HRP immobilized on PEGMA microgels. These experiments show that PEGMA microgels can effectively stabilize the enzymes like HRP against thermal denaturation. Nevertheless, if we compare the value of kdecay‑PEGMA obtained in this study with the value of the decay rate of HRP immobilized on PHEAA microgels from our previous study,27 kdecay‑PHEAA = 6.5 × 10−4 min−1, the PEGMA microgels appear be somewhat less effective than PHEAA in stabilizing HRP enzymes against thermal denaturation.

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CONCLUSIONS We report the synthesis and characterization of dual responsive PEGMA-MAA microgels composed of temperature-responsive OEGMA300 and pH-responsive MAA moieties. PEGMA-MAA microgels exhibited a thermal VPPT at 60 °C in DI water and in acidic buffers but remained unresponsive in neutral and basic buffers. In spite of the high −COOH group content (31 mol % MAA groups), these microgels showed a strong resistance against the covalent immobilization of enzymes to the backbone MAA groups. To overcome this problem, we I


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Macromolecules attached a diamine spacer with four EG units to the −COOH groups of the microgel. In this way, we were able to attach the enzyme HRP to the microgel using BAH chemistry. We found that over the temperature range of 25−50 °C the PEGMA-HRP was about a factor of 3 less active than the native enzyme. At higher temperatures, the enzyme activity of both the immobilized enzyme and native HRP decreased compared to the maximum activity at ca. 50 °C. The extent of decrease for the native enzyme was more pronounced compared to that of the microgel-bound enzyme, and there was no obvious change in activity associated with the VPPT of the microgel. Immobilization of HRP on the PEGMA-MAA microgels also stabilized HRP against thermally induced denaturation during prolonged annealing at 50 °C. While PHEAA microgels are somewhat more effective than PEGMA-MAA microgels at providing enhanced stability to HRP, PEGMA-MAA microgels are easier to synthesize. Our strategy for bioconjugation of PEGMA-MAA could be of potential interest for a broader application of PEG-based microgels in biotechnology.

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(7) Gan, T.; Zhang, Y.; Guan, Y. In Situ Gelation of P(NIPAMHEMA) Microgel Dispersion and Its Applications as Injectable 3D Cell Scaffold. Biomacromolecules 2009, 10, 1410−1415. (8) Madsen, J.; Armes, S. P. (Meth)acrylic Stimulus-Responsive Block Copolymer Hydrogels. Soft Matter 2011, 8, 592−605. (9) Lutz, J.-F.; Weichenhan, K.; Akdemir, Ö .; Hoth, A. About the Phase Transitions in Aqueous Solutions of Thermoresponsive Copolymers and Hydrogels Based on 2-(2-Methoxyethoxy)ethyl Methacrylate and Oligo(ethylene Glycol) Methacrylate. Macromolecules 2007, 40, 2503−2508. (10) Lutz, J.-F. Thermo-Switchable Materials Prepared Using the OEGMA-Platform. Adv. Mater. 2011, 23, 2237−2243. (11) Lutz, J.-F.; Akdemir, Ö .; Hoth, A. Point by Point Comparison of Two Thermosensitive Polymers Exhibiting a Similar LCST: Is the Age of Poly(NIPAM) Over? J. Am. Chem. Soc. 2006, 128, 13046−13047. (12) Lutz, J.-F. Polymerization of Oligo(ethylene Glycol) (Meth)acrylates: Toward New Generations of Smart Biocompatible Materials. J. Polym. Sci., Part A: Polym. Chem. 2008, 46, 3459−3470. (13) Lutz, J.-F.; Hoth, A. Preparation of Ideal PEG Analogues with a Tunable Thermosensitivity by Controlled Radical Copolymerization of 2-(2-Methoxyethoxy)ethyl Methacrylate and Oligo(ethylene Glycol) Methacrylate. Macromolecules 2006, 39, 893−896. (14) Knop, K.; Hoogenboom, R.; Fischer, D.; Schubert, U. S. Poly(ethylene Glycol) in Drug Delivery: Pros and Cons as Well as Potential Alternatives. Angew. Chem., Int. Ed. 2010, 49, 6288−6308. (15) Cai, T.; Marquez, M.; Hu, Z. Monodisperse Thermoresponsive Microgels of Poly(ethylene Glycol) Analogue-Based Biopolymers. Langmuir 2007, 23, 8663−8666. (16) Lutz, J.-F.; Andrieu, J.; Ü zgün, S.; Rudolph, C.; Agarwal, S. Biocompatible, Thermoresponsive, and Biodegradable: Simple Preparation of “All-in-One” Biorelevant Polymers. Macromolecules 2007, 40, 8540−8543. (17) Lee, H.; Lee, E.; Kim, D. K.; Jang, N. K.; Jeong, Y. Y.; Jon, S. Antibiofouling Polymer-Coated Superparamagnetic Iron Oxide Nanoparticles as Potential Magnetic Resonance Contrast Agents for in Vivo Cancer Imaging. J. Am. Chem. Soc. 2006, 128, 7383−7389. (18) Lutz, J.-F.; Stiller, S.; Hoth, A.; Kaufner, L.; Pison, U.; Cartier, R. One-Pot Synthesis of PEGylated Ultrasmall Iron-Oxide Nanoparticles and Their in Vivo Evaluation as Magnetic Resonance Imaging Contrast Agents. Biomacromolecules 2006, 7, 3132−3138. (19) Hu, Z.; Cai, T.; Chi, C. Thermoresponsive Oligo(ethylene Glycol)-Methacrylate- Based Polymers and Microgels. Soft Matter 2010, 6, 2115. (20) Lee, B. S.; Lee, J. K.; Kim, W.-J.; Jung, Y. H.; Sim, S. J.; Lee, J.; Choi, I. S. Surface-Initiated, Atom Transfer Radical Polymerization of Oligo(ethylene Glycol) Methyl Ether Methacrylate and Subsequent Click Chemistry for Bioconjugation. Biomacromolecules 2007, 8, 744− 749. (21) Trmcic-Cvitas, J.; Hasan, E.; Ramstedt, M.; Li, X.; Cooper, M. A.; Abell, C.; Huck, W. T. S.; Gautrot, J. E. Biofunctionalized Protein Resistant Oligo(ethylene Glycol)-Derived Polymer Brushes as Selective Immobilization and Sensing Platforms. Biomacromolecules 2009, 10, 2885−2894. (22) Lee, B. S.; Chi, Y. S.; Lee, K.-B.; Kim, Y.-G.; Choi, I. S. Functionalization of Poly(oligo(ethylene Glycol) Methacrylate) Films on Gold and Si/SiO 2 for Immobilization of Proteins and Cells: SPR and QCM Studies. Biomacromolecules 2007, 8, 3922−3929. (23) Roth, P. J.; Jochum, F. D.; Zentel, R.; Theato, P. Synthesis of Hetero-Telechelic A,ω Bio-Functionalized Polymers. Biomacromolecules 2010, 11, 238−244. (24) Chen, G.; Huynh, D.; Felgner, P. L.; Guan, Z. Tandem Chain Walking Polymerization and Atom Transfer Radical Polymerization for Efficient Synthesis of Dendritic Nanoparticles for Bioconjugation. J. Am. Chem. Soc. 2006, 128, 4298−4302. (25) Hucknall, A.; Kim, D.-H.; Rangarajan, S.; Hill, R. T.; Reichert, W. M.; Chilkoti, A. Simple Fabrication of Antibody Microarrays on Nonfouling Polymer Brushes with Femtomolar Sensitivity for Protein Analytes in Serum and Blood. Adv. Mater. 2009, 21, 1968−1971.

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.macromol.6b01270. Additional results and discussion, TEM images, DLS CONTIN plots, titration plots, LCFM images and enzyme kinetic plots, and a table comparing conversion efficiencies of PEGMA-MAA microgels with PHEAA microgels (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail [email protected] (M.A.W.). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors thank the Natural Sciences and Engineering Research Council (NSERC) Canada for their support of this work.

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REFERENCES

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PEGMA-Based Microgels: A Thermoresponsive Support for Enzyme

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