Temperature-Invariant Aqueous Microgels as Hosts for

Article pubs.acs.org/Biomac

Temperature-Invariant Aqueous Microgels as Hosts for Biomacromolecules Sepehr Mastour Tehrani,†,‡ Yijie Lu,‡ Gerald Guerin,‡ Mohsen Soleimani,†,‡ Dmitry Pichugin,‡ and Mitchell A. Winnik*,†,‡ †

Downloaded by UNIV OF PRINCE EDWARD ISLAND on September 8, 2015 | http://pubs.acs.org Publication Date (Web): September 3, 2015 | doi: 10.1021/acs.biomac.5b00768

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: Immobilization of enzymes on solid supports has been widely used to improve enzyme recycling, enzyme stability, and performance. We are interested in using aqueous microgels (colloidal hydrogels) as carriers for enzymes used in high-temperature reactions. These microgels should maintain their volume and colloidal stability in aqueous media up to 100 °C to serve as thermo-stable supports for enzymes. For this purpose, we prepared poly(Nhydroxyethyl acrylamide) (PHEAA) microgels via a two-step synthesis. First, we used precipitation polymerization in water to synthesize colloidal poly(diethylene glycol-ethyl ether acrylate) (PDEGAC) particles as a precursor. PDEGAC forms solvent swollen microgels in organic solvents such as methanol and dioxane and in water at temperatures below 15 °C. In the second step, these PDEGAC particles were transformed to PHEAA microgels through aminolysis in dioxane with ethanolamine and a small amount of ethylenediamine. Dynamic laser scattering studies confirmed that the colloidal stability of microgels was maintained during the aminolysis in dioxane and subsequent transfer to water. Characterization of the PHEAA microgels indicated about 9 mol % of primary amino groups. These provide functionality for bioconjugation. As proof-of-concept experiments, we attached the enzyme horseradish peroxidase (HRP) to these aqueous microgels through (i) N-(3(dimethylamino)propyl)-N′-ethylcarbodiimide hydrochloride (EDC) coupling to the carboxylated microgels or (ii) bis-aryl hydrazone (BAH) coupling to microgels functionalized with 6-hydrazinonicotinate acetone (PHEAA-HyNic). Our results showed that HRP maintained its catalytic activity after covalent attachment (87% for EDC coupling, 96% for BAH coupling). The microgel enhanced the stability of the enzyme to thermal denaturation. For example, the residual activity of the microgelsupported enzyme was 76% after 330 min of annealing at 50 °C, compared to only 20% for the free enzyme under these conditions. PHEAA microgels in water show great promise as hosts for enzymatic reaction, especially at elevated temperatures.

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INTRODUCTION Enzymes have numerous applications in industrial processes due to their high specificity, stereoselectivity, and catalytic efficiency. In line with the new developments in enzymology, there is an ongoing interest in enzyme immobilization on particle supports. Immobilization can facilitate enzyme recovery and often improve stability.1−3 In addition, immobilization can circumvent enzyme aggregation, which is typical of native enzymes in solution.4 When enzymes are immobilized on a particle support, especially through a multipoint covalent attachment, enzyme stability is increased. This added stability of immobilized enzyme is particularly useful for applications involving extensive changes of temperature or pH.5−8 Some research groups have also shown that enzyme stability is improved with the entrapment of enzymes in the pores of hydrogels or silica.9−12 However, covalent attachment of enzymes to particle supports is often preferred over other immobilization techniques because it avoids possible enzyme leaching into the surrounding medium.13 © XXXX American Chemical Society

For enzymatic reactions, different types of bead supports have been investigated. Typical supports for enzyme immobilization are porous glass beads, silica beads, polystyrene (PS) beads, copolymers of acrylamide, copolymers of (meth)acrylates, agarose, and chitosan.4,14−16 These beads are normally large (tens of μm to mm in diameter), and thus they are not colloidal. One prominent industrial example of colloidal supports is those used in DNA amplification with polymerase chain reaction (PCR).17 In this process, (1−5 μm) cross-linked PS microbeads are coupled with DNA primers, suspended in water-in-oil emulsions (ePCR), and subjected to thermocycling (e.g., from 45−92 °C). Cross-linked PS microbeads are also used in bead-assays and in multiplexed protein detection technologies.18,19 Although PS microbeads have found numerous applications, there are some drawbacks Received: June 8, 2015 Revised: July 24, 2015

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DOI: 10.1021/acs.biomac.5b00768 Biomacromolecules XXXX, XXX, XXX−XXX

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Biomacromolecules

these hydrophilic and water-swellable colloidal particles as supports for immobilization of antibodies and enzymes with applications in PCR. The prerequisites for the microgel supports used for hightemperature processes are to maintain their size, colloidal stability, and chemical stability. Such supports could also be useful in applications such as microreactors or catalyst carriers, which require support stability in severe conditions.42,43 Since PNIPAm and most aqueous microgels are temperature sensitive and shrink upon heating, high-temperature enzymatic reactions are rarely examined on these hydrogel particles.29,44 In systems with enzymes immobilized on aqueous microgels at temperatures close to the VPTT of the microgels, the network starts to lose its water content and the microgels collapse. The dehydration and shrinkage can severely suppress substrate and product mass transfer and “shut-down” the catalytic activity of enzymes.45 Also, the increased hydrophobic interactions above the VPTT of the support can negatively impact the native functionality of most enzymes and proteins.46,47 There are a few exceptions, for example, enzymes that operate effectively in a hydrophobic environment and maintain their activity above the VPTT of the PNIPAm microgel support.44 We anticipate that aqueous microgels that do not have a VPTT would be a more versatile support for enzymatic reactions at higher temperatures. Despite the large number of research groups investigating thermosensitive microgels, there is almost no research on temperature-invariant aqueous microgels. From this perspective, we are interested in the preparation of aqueous microgels based on poly(N-hydroxyethyl acrylamide) (PHEAA). This polymer has the highest water solubility among nonionic polymers.48 Albarghouthi et al.49 suggested that PHEAA is even more hydrophilic than commonly known hydrophilic polymers such as poly(dimethylacrylamide) and polyacrylamide. Some research groups have been interested in taking advantage of this high level of polarity and hydrophilicity to reduce nonspecific adsorption of proteins. For example, Zhao and Zheng50 showed that this polymer has almost zero nonspecific adsorption in long-term exposure to blood plasma or serum. They also showed that PHEAA coated surfaces have exceptional antifouling properties against bacterial growth. This group prepared nanogels based on PHEAA with an average diameter of 130−160 nm through a process employing inverse microemulsion polymerization.51,52 There are no reports suggesting that PHEAA has an LCST below 100 °C. In addition, it has been shown that this polymer has good resistance against thermal degradation and hydrolysis.53,54 As a consequence, PHEAA microgels are an interesting candidate as a support for protein analysis and enzyme immobilization for applications in aqueous media at high temperatures. In our design, we anticipate that aqueous microgels of PHEAA will have three important benefits. First, the high water content should provide a well-hydrated environment for covalently attached biomacromolecules. Second, the porosity associated with the colloidal hydrogels should increase the number of available sites for biomacromolecule attachment or hybridization. Finally, the absence of a VPTT below 100 °C and the hydrolytic stability of this polymer will make these microgels particularly suitable as hosts for high-temperature enzymatic reactions in water. In this study, we developed a two-stage approach for the synthesis of PHEAA microgels. In the first step, we used precipitation polymerization55 to prepare cross-linked precursor particles of poly(diethylene glycol-ethyl ether acrylate)

associated with using these microbeads in protein analysis. The hydrophobicity of the PS matrix and nonspecific adsorption of proteins on it are among the problems that have raised interest in searching for replacements for PS microbeads. Particles chosen as a carrier for an enzyme or a bioconjugation agent for multiplexed assays should, ideally, have no nonspecific interactions with biomacromolecules.20,21 Nonspecific adsorption of DNA or enzymes on hydrophobic surfaces can distort their shape and interfere with analytical measurements. Hydrophilic matrices are preferred because they tend to have much lower levels of nonspecific interactions with the biologically active macromolecules. Hydrogels are accepted as one of the most promising carriers for biomacromolecules because they carry great amounts of water.22,23 When proteins and enzyme are maintained in wet surroundings as in hydrogels, they can fold properly and demonstrate their full biological functionality, whereas on a hydrophobic surface, proteins can unfold, spread over the support, and lose their native properties.24 Different research groups have used this concept and designed a variety of hydrogel hosts for biomacromolecule attachment.25,26 For example, Doyle and his group27 used hydrogel microparticles of poly(ethylene glycol) and poly(ethylene glycol diacrylate) (PEGDA) prepared in a microfluidic channel as a support for microRNA profiling and for performing rolling circle amplification (RCA) reactions. Pelton, Li, and colleagues investigated another example of biomacromolecule immobilization on aqueous microgels (colloidal hydrogels) of poly(N-isopropylacrylamide) (PNIPAm).28 They demonstrated that oligoDNA-PNIPAm microgel conjugates were compatible with enzymatic reactions used for amplification of DNA at room temperature. PNIPAm copolymer microgels have also been used for immobilization of water-soluble enzymes for use in organic media.29−31 PNIPAm copolymer microgels and bulk hydrogels are thermosensitive, displaying a volume phase transition temperature (VPTT) between 30 and about 45 °C depending on composition. Exceeding this temperature, the so-called lower critical solution temperature (LCST) of the polymer, the hydrophobic nature of the gel dominates and collapses the gel.32,33 This change in microgel properties at elevated temperatures can interfere with their use as supports for enzyme catalyzed reactions. Performing enzymatic reaction at high temperature offers several advantages. For instance, at elevated temperatures, the solubility of reactants and mass transfer rates improve. Moreover, high temperature can diminish the effect of impurities and suppress the influence of harmful bacteria that may contaminate the enzymes.34 For some applications, such as DNA amplification using PCR, a high-temperature step (∼92 °C) is employed to melt DNA double strands at each cycle of amplification. In RCA amplification of DNA, high temperature is also preferred for the detection of diseased genes associated with single nucleotide polymorphism (SNPs) because the elevated temperature provides greater discrimination between fully matched and single-base mismatched DNA targets.35 Recently, Schierack and coauthors17 reviewed different physical and chemical requirements for microbeads used in PCR. Their bead-assay platform for protein and nucleic acid analysis employs “thermo-tolerant” poly(methyl methacrylate) (PMMA) microbeads.36,37 Horak and colleagues38−41 prepared a series of poly(hydroxyethyl methacrylate) (PHEMA)-based microbeads with diameters ranging from 1−3 μm and used B


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Biomacromolecules

prepared by placing one drop of a microgel solution in phosphate buffer pH 7.2 between a microscopy slide and a coverslip. Dynamic Laser Scattering (DLS). DLS measurements were performed on a Malvern Zetasizer Nano ZS instrument at a backscattering angle of 173°. For measurements involving temperature change, Zetasizer folded capillary cells were used, and samples were allowed to equilibrate for 10 min at each temperature. Nuclear Magnetic Resonance (NMR). 1H NMR spectra of small molecules and soluble polymers were recorded either on Varian Mercury plus NMR spectrometer operating at 400 MHz or Agilent DD2−500 operating at 500 MHz equipped with a Xsens cryo-probe. For microgel samples, proton high-resolution NMR spectra were measured under magic-angle sample spinning (MAS) conditions on an Agilent DD2−700 NMR spectrometer operating at 700 MHz with a 4 mm FASTnano probe. The measurements were performed at rotor frequencies of 4 kHz and 8 kHz. MestReNova software was used for the spectra processing. 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. Materials. Monomer di(ethylene glycol) ethyl ether acrylate (DEGAC), initiator ammonium persulfate (APS, 98%), cross-linker N,N′-methylene bis(acrylamide) (BIS, 99%), ethylenediamine (>99%), ethanolamine (>99%), 1,4-dioxane (anhydrous, 99.8%), picrylsulfonic acid solution (5% w/v in H2O, TNBS assay), sodium dodecyl sulfate (SDS, 99%), hydrochloric acid and sodium hydroxide (1N, HPLC standard), fluorescein isothiocyanate (FITC, 98% HPLC), peroxidase from horseradish (HRP, type VI-A), 2,2′-Azino-bis(3ethylbenzothiazoline-6-sulfonic acid) diammonium salt (ABTS assay, 98% HPLC), hydrogen peroxide (30% wt. in H 2O), N-(3(dimethylamino)propyl)-N′-ethylcarbodiimide hydrochloride (EDC, 98%), N-hydroxysuccinimide (NHS, 98%), Bradford reagent (Coomassie dye protein assay), 4-formylbenzoic acid (97%), 6-chloropyridine-3-carboxylic acid (99%), hydrazine hydrate, anhydrous DMF (99.8%), and trimethylamine (99%) were purchased from SigmaAldrich. For preparing buffer solutions, sodium bicarbonate, sodium phosphate monobasic, potassium phosphate monobasic, phosphate buffered saline (PBS, tablet), sodium acetate, and 2-(N-morpholino)ethanesulfonic acid hydrate (MES hydrate) were prepared in MiliQ water (gradient filter, Milipore). Succinimidyl 4-formylbenzoate (S-4FB) and succinimidyl 6hydrazinonicotinate (S-HyNic) were synthesized as described by Grotzky et al.59 PDEGAC Particle Synthesis. Precipitation polymerization of DEGAC 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. DEGAC monomer (2 g, 10 mmol) and BIS cross-linker (0.10 g, 0.65 mmol) were dissolved in MiliQ water (95 mL) and transferred to the flask. The mixture was heated to 70 °C and bubbled with nitrogen for 45 min, and then APS initiator (21.7 mg, 95 μmol) dissolved in MiliQ 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, the homogeneous white microgel solution obtained was transferred to centrifugation tubes for purification. The microgel solution was washed three times by sedimentation and redispersion in cold water (4 °C), well below the VPTT of PDEGAC (15 °C), using a Sorvall Evolution RC superspeed centrifuge at 20 500 rpm for 20 min to remove all side products. After sample withdrawal for characterization purposes, the remaining microgel mixture was solvent exchanged first to methanol via two sedimentation−redispersion cycles and then to 1,4-dioxane via an additional three cycles of sedimentation and redispersion. After the final cycle, the microgel sample in anhydrous dioxane was sealed and stored at room temperature. Aminolysis. For the aminolysis step, aliquots of the PDEGAC microgel (160 mg) in dioxane (0.8 mL), ethanolamine (1.2 mL, ∼ 20 mmol), and ethylenediamine (60 μL, ∼ 0.9 mmol) were added in a single-necked round-bottom flask and mixed with a Teflon-coated

(PDEGAC). Note that this material is also a microgel. This polymer has an LCST in water of about 15 °C,56 and thus these particles are collapsed in water both at the synthesis temperature (70 °C) and at room temperature. In the second step, these particles were subjected to aminolysis in dioxane with a mixture of ethanolamine and ethylene diamine to convert the PDEGAC microgels in dioxane to amine-functional PHEAA-NH2 microgels. The synthetic design is presented in Scheme 1. After transfer to aqueous solution, the PHEAA


Scheme 1. Multi-step Preparation of PHEAA Microgels Loaded with HRP Enzyme

microgels assumed their water-swollen dimensions. Our results show that these microgels in water preserved their size and colloidal stability when heated, even to high temperatures. To test the effectiveness of our PHEAA microgels as a support for enzyme reactions, we chose horseradish peroxidase (HRP) as a model enzyme. HRP is a 44 kDa glycoprotein with six lysine residues that is widely studied and often conjugated to a labeled molecule.57,58 After the covalent attachment of HRP to the PHEAA microgels, the amount of enzyme loaded in the microgels was determined, and the influence of attachment to the microgels on the kinetic and thermal stability of the enzyme was examined in phosphate buffer. Kinetic studies showed that immobilization on these PHEAA microgels did not interfere with the enzymatic activity of HRP and, more importantly, led to improved enzyme stability and activity at high temperatures compared to the native enzyme. These microgels show great promise as supports for a wide variety of enzyme reactions in water.

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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 100 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. Samples were diluted with DI water and deposited on either an ultrathin carbon-coated copper grid or a PELCO graphene TEM support film on Lacey carbon. Laser Scanning Confocal Fluorescence Microscopy (LCFM). LCFM images were obtained using a Leica TCS SP2 confocal microscope equipped with a 63× oil objective, numerical aperture (NA) = 1.4, with excitation wavelength λex = 488 nm and emission wavelength of λem = 600 ± 25 nm. The samples for LCFM were C


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Biomacromolecules magnetic stir bar. At room temperature, the mixture was bubbled with nitrogen for 1 h and then immersed in a silicone oil bath at 75 °C. The reaction was stirred at this temperature for 24 h. The resulting mixture turned transparent at the end of the reaction. After the reaction, we used two cycles of sedimentation and redispersion in methanol followed by dilution of product with acetic acid (20% w/v). The product was further purified through five cycles (45 min each) of spin filtration with 10k MWCO spin filters using DI water as diluent. At the end, product microgels were dispersed in DI water and stored at room temperature to serve as a stock mixture. The final concentration of microgels in water was 23 mg/mL. Attempted Aminolysis in Methanol. The PDEGAC microgels (160 mg) were dispersed in methanol (0.8 mL) and mixed with ethylenediamine (1.2 mL, ∼18 mmol) in a single-necked roundbottom flask under magnetic mixing. At room temperature, the mixture was bubbled with nitrogen for 1 h and then immersed in 59 °C silicon bath, and the reaction was let to continue for 24 h. After the reaction, the product was diluted with acetic acid (20% w/v) and subjected to several cycles of purification with 10k MWCO spin filters using DI water. The product of this reaction is a poly(aminoethyl acrylamide) (PAMEA) microgel. Free Amine Quantification by TNBS Assay. The TNBS assay was used to quantify the primary amine content of PHEAA-NH2 microgels. The procedure was performed as described by Hermanson.60 Briefly, TNBS was dissolved in sodium bicarbonate buffer, (100 mM, pH 8.5) (TNBS assay buffer) at a concentration of 0.01% (w/v). PHEAA-NH2 microgels were also resuspended in the assay buffer at a concentration of 200 μg/mL. An aliquot of the microgel mixture (1.0 mL) was added to the TNBS solution (0.5 mL), and the mixture was incubated 2 h at 37 °C. Then an SDS solution (1 mL, 10% w/v) and HCl (0.5 mL, 1 N) were added. The UV−vis absorbance of the solution was measured at 420 nm, at room temperature. Standard curves were plotted from assaying different concentrations of ethanolamine after treatment with the TNBS solution. These calibration curves are shown in Figure S6. FITC Labeling. To label PHEAA-NH2 microgel with a fluorescent dye, FITC (1 mg, 2.5 μmol) was dissolved in DMSO (100 μL). An aliquot of PHEAA stock solution (50 μL, 1.15 mg microgel) was dispersed in NaHCO3 buffer (450 μL, 100 mM, pH 8.5). The microgel and FITC solutions were mixed together in an amber vial under magnetic stirring. The reaction was continued for 6 h at room temperature, and then the mixture was washed extensively with phosphate buffer (100 mM, pH 7.2) using a 10k MWCO spin filter until the filtrate became colorless. For confocal microscopy analysis, the sample was diluted visually so that it had a pale green color. Succinic Anhydride (SA) Reaction. To decorate the PHEAA-NH2 microgels with COO− functionality, 1.5 mL of PHEAA-NH2 stock solution (34.5 mg PHEAA-NH2 microgel) was buffer exchanged through spin filtering with sodium bicarbonate buffer (100 mM, pH 8.5) and finally redispersed in 1 mL of this buffer. To this mixture, solid SA (300 mg, 3 mmol) was added at room temperature. During the first hour of the reaction, to keep the pH of the mixture basic, several aliquots of NaOH solution (0.1 N, 100 μL) were added. To make sure that all of the amine groups had reacted, 2 h after the start of the reaction, additional SA (200 mg) was added, and the reaction was allowed to proceed overnight. After the reaction, the PHEAA-SA microgels were purified with DI water and phosphate buffer pH 7.2 using a 10k MWCO spin filter. EDC Coupling of PHEAA-SA to HRP. The strategy applied for HRP conjugation was modified slightly from the method described by Hermanson.60 In our work, PHEAA-SA microgels (3 mg) were suspended in MES buffer (250 μL, 50 mM, pH 6.0), mixed with EDC (100 mg) in a 2 mL Eppendorf tube, and allowed to stand for 30 min. Then the solution was washed quickly with 10k MWCO spin filters using MES buffer (50 mM, pH 6.0) to strip off extra EDC. The EDCactivated PHEAA-SA was then resuspended in MES buffer (200 μL) and incubated in 1.5 mL tube containing the HRP enzyme (1 mg), and the incubation was continued overnight. After the reaction, the microgels conjugated to HRP were separated from the unreacted HRP via repeated cycles of centrifugation at 20 500 rpm with phosphate buffer (50 mM, pH 6.8). Washing cycles continued until no enzymatic

activity could be observed from the supernatant. PHEAA-EDC microgels were resuspended in the phosphate buffer pH 6.8 and stored at 4 °C. PHEAA-NH2 Microgel Coupling to HyNic. PHEAA-NH2 microgels (4 mg) were suspended in phosphate buffer (1.5 mL, 100 mM, pH 8.0). To this solution, an aliquot of HyNic (1.01 mg, 3.5 μmol) in DMSO (350 μL) was added. The reaction continued for 3 h at room temperature, and then unreacted HyNic was washed off the microgel mixture by repeated spin filtering using a 3000 MWCO membrane and acetate buffer (100 mM, pH 5.0). The microgels were resuspended in this acetate buffer and diluted to a volume of 290 μL. An aliquot of this solution (20 μL) was withdrawn to quantify the number of HyNic per microgel, and the remaining 270 μL was reacted with the S-4FB modified HRP (see the following). To evaluate the number of HyNic per microgel, 4-formylbenzoic 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). HRP Coupling to S-4FB. HRP enzyme (0.9 mg, ca. 20 nmol) was dissolved in phosphate buffer (250 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.59 The reaction continued for 3 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 acetate 100 mM buffer pH 5.0. After extensive washing cycles, 4FB-modified HRP was suspended in acetate buffer (114 μL, 100 mM, pH 5.0). An aliquot of this solution (20 μL) was withdrawn to quantify the number of 4FB per HRP molecule, and the remaining 94 μL was reacted with the HyNic-decorated PHEAA microgel. To evaluate the number of 4FB groups per HRP molecule, a solution of 2-hydrazinopyridine (80 μL, 500 μM) in acetate 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 = 18 000 M−1 cm−1). PHEAA-HyNic and HRP-4FB Conjugation. The PHEAA-HyNic solution (270 μL) and the HRP-4FB solution (94 μL) (described earlier) in acetate buffer (pH 5.0) 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 (4 h). At the end of the reaction, nonconjugated HRP-4FB was separated from conjugated PHEAA-HRP by centrifugation at 20 500 rpm and resuspension in phosphate buffer pH 6.8. This sample is denoted PHEAA-BAH and was stored at 4 °C. Enzyme Loading Quantification (Bradford Assay). The content of enzyme loaded into the PHEAA microgels was determined from a Bradford microassay. In this method, Bradford reagent (200 μL) was incubated with the enzyme-loaded microgel sample of known microgel concentration suspended in phosphate buffer (50 mM, pH 5.2). Samples were incubated at room temperature, and an absorbance measurement at 595 nm was taken 15 min after the start of the incubation. Two standard curves were plotted for an accurate quantification. First, known amounts of unloaded PHEAA microgel were assayed against Bradford reagent to determine the net contribution of microgel−dye interaction in the assay. This type of calibration was reported by Gawlitza et al. for PNIPAm microgels.61 The second standard curve was prepared by assaying HRP at different concentrations (0−50 μg/mL) against Bradford reagent. Calibration curves are shown in Figure S7. 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 of the ABTS assay at varying concentrations of the ABTS substrate. To perform this assay, an aliquot of HRP solution (50 μL, 1.76 μg/mL, 40 nM) in phosphate buffer (50 mM, pH 6.8), H2O2 (50 μL, 0.3 wt %, 88 mM), and varying amounts of ABTS solution (0−500 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 measured by recording the change in the absorbance of the solution at 405 nm D


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Biomacromolecules 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 following equation:


ϑ=

Vmax[s] K m + [s]

(NAD) polymerization is in principle possible. This is a precipitation polymerization in a nonaqueous solvent in which the monomer, initiator, and other reactants are soluble in the medium, but the polymer precipitates as it is formed. Horak and colleagues63 have reported the synthesis of particles consisting of linear poly(N,N-dimethyl acrylamide) by NAD polymerization. Synthesizing cross-linked particles by dispersion polymerization poses another level of difficulty. All attempts on our part to synthesize PHEAA microgels by NAD polymerization met with failure. One can also consider microemulsion polymerization. It can be useful for synthesizing small particles. We propose an alternative two-stage approach involving precursor particles that could be converted to PHEAA through a chemical modification. For this purpose, we chose precipitation polymerization in water of DEGAC. PDEGAC has a LCST around 15 °C. DEGAC polymerization was carried out at 70 °C in water with a persulfate initiator and 6 mol % BIS cross-linker and continued for 18 h. DLS measurements of this PDEGAC dispersion at 25 °C in pH 7.2 buffer gave a Zaverage diameter dZ = 450 nm (see Table 1). A SEM image of

(1)

where [s] is the ABTS substrate concentration, and ϑ is the initial rate. Microsoft Excel was used for curve fitting. Data fitted to eq 1 are plotted in Figure S8. Activity Measurement 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), phosphate buffer (400 μL, pH 5.2, 50 mM), and 50 μL of (1.76 μg/mL) native or immobilized HRP in phosphate buffer (pH 6.8, 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, the 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 (1.76 μg HRP/mL) 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 (30 min annealing) as t = 0:

(%) residual activity = α exp(− kdecay × t )

Table 1. DLS Measurements of PDEGAC and PHEAA-NH2 Microgel Diameters at 25 °C for Samples Dispersed in 100 mM Phosphate Buffer pH 7.2 and in Anhydrous Dioxane microgel

aqueous buffer, 25 °C dZ, nm, (PDI)a

dioxane, 25 °C dZ, nm, (PDI)a

PDEGAC PHEAA-NH2

449 (0.03) 817 (0.03)

503 (0.02) 913 (0.26)b

a Values of dz were calculated by a cumulant analysis of DLS autocorrelation decay data. CONTIN plots and autocorrelation decay profiles are presented in Figure S3. bAlthough the CONTIN plot for this sample was sharp and monomodal, a significantly higher polydispersity was calculated by the cumulant analysis, and one can see a broader decay of the autocorrelation function for this sample (Figure S3E).

(2)

where α is a pre-factor.

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RESULTS AND DISCUSSION Microgel Preparation. The main purpose of this research was to synthesize aqueous microgels that do not exhibit a temperature response below 100 °C in water. By design, the polymer backbone of the microgel should not have an LCST up to this temperature. Most hydrophilic polymers demonstrate LCST behavior in aqueous solution. At elevated temperatures, hydrogen bonds between the polymer and water molecules become weak, and the polymer starts to precipitate. There are few choices of hydrophilic polymers without a temperature response in water. Polyacrylamide, poly(N,N-dimethyl acrylamide) (PDMA), and PHEAA are polymers without a reported LCST in water. PHEAA is particularly attractive since the hydroxyl in the pendant group renders the polymer particularly hydrophilic compared to other nonionic polymers. While linear hydrophilic polymers potentially lacking an LCST like PHEAA are readily synthesized by free radical polymerization, it is a challenge to synthesize them as microgels.62 The conventional method for preparing hydrophilic microgels is precipitation polymerization in water. In such systems, the ingredients, including the monomers, crosslinker, and initiator, are dissolved in water, and at the polymerization temperature (e.g., 50−70 °C), the propagating polymer chains become insoluble and start to precipitate. The precipitates serve as nuclei for further microgel growth. Since polymers without an LCST remain soluble in water at all accessible temperatures, precipitation polymerization of these polymers in water is not possible. Nonaqueous dispersion

these particles is presented in Figure 1, panel A, and another SEM image, taken with a tilted grid, is shown in Figure S1 (Supporting Information). This latter image emphasizes that these soft particles form flattened structures on the grid. From the histogram of particle diameters in Figure 1, panel B, we calculate a number average diameter dn = 440 nm. The polymerization reaction described earlier also works well with lower levels of BIS cross-linker, but the particles obtained are more difficult to centrifuge, particularly at later stages of the chemical transformation. We used multiple sedimentation− redispersion cycles for sample isolation and purification. Thus, for these proof-of-concept experiments, we chose to work with the particles prepared with 6 mol % BIS cross-linker. As we will see, this high level of cross-linker likely limits the number of enzymes that can be attached to the PHEAA microgels obtained after the chemical transformations. To convert PDEGAC to PHEAA, the PDEGAC particles were first transferred to dioxane and then subjected to exhaustive aminolysis with an excess of ethanolamine plus ethylenediamine (22:1 mol ratio). This reaction is shown in Figure 2, panel A. During the transfer from water to dioxane through multiple sedimentation and redispersion steps, the initially turbid dispersion of PDEGAC particles became much less cloudy but retained an obvious haziness that increased noticeably when the ethanolamine and ethylenediamine were E


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Biomacromolecules

Figure 1. (A) SEM image of PDEGAC microgels obtained from precipitation polymerization at 70 °C. (B) Histogram analysis of PDEGAC microgels SEM images obtained from measuring at least 100 individual microgels using ImageJ software.

A 1H NMR spectrum of the PDEGAC microgels dispersed in methanol-d4 is presented in Figure 2, panel B. The sharp peaks are a signature of highly mobile pendant groups present in the solvent swollen microgel network. To assess the effectiveness of the aminolysis reaction, we paid particular attention to the peak at 4.2 ppm associated with the acrylate ester CH2 group. After the aminolysis reaction, attempts to obtain traditional 1H NMR spectra in methanol-d4 or D2O led to very broad peaks. As a consequence, we turned to a high-resolution magic angle spinning (HR-MAS) probe to obtain the spectrum in D2O, as shown in Figure 2, panel C. The most obvious feature of this spectrum is the disappearance of the 4.2 ppm peak, indicating complete reaction of the acrylate ester groups. The inset in Figure 2, panel C provides information about the −CH2NH2 group content of the PHEAA microgel. This aspect will be discussed in a later section of this paper. One further comment is in order about the reaction. We found that multiple exchanges of the PDEGAC particle suspension with anhydrous dioxane were necessary to avoid unwanted hydrolysis of the acrylate ester groups to acrylic acid. When we ran the aminolysis reaction in methanol instead of anhydrous dioxane, we found incomplete conversion and evidence by 1H NMR about 15 mol % carboxylic acid groups in the product (Figure S2). Particle Size Changes Accompanying the Chemical Transformation. A TEM image of the PHEAA-NH2 microgels is shown in Figure 3, panel A. The microgels appeared to form hairy globular structures on the TEM grid. The histogram analysis of microgel size distribution is shown in Figure 3, panel B. The average diameter of PHEAA-NH2 microgels on TEM grid is about 240 nm. By comparing the TEM image of the PHEAA-NH2 microgels in Figure 3 with the SEM image of the precursor particles in Figure 1, one can see that the number average diameter of the PHEAA-NH2 microgels (240 nm) was significantly smaller than that (440 nm) of the precursor particles, although the size distribution was maintained. The smaller size is likely a consequence of the mass loss during the chemical transformation. Both the SEM and TEM images were obtained for samples in the dry state, in which the microgels are dehydrated and collapsed. For the hydrated PHEAA-NH2 microgels in phosphate buffer at pH 7.2, we found dZ = 817 nm (Table 1), almost two-fold larger than that of its precursor PDEGAC particles in water (dZ = 449 nm). This increase in size originates from the enhanced hydrophilicity of the

Figure 2. (A) The aminolysis reaction of PDEGAC particles in dioxane with excess ethanolamine plus ethylenediamine (22:1 mol ratio). (B) 1H NMR spectra of PDEGAC microgels in methanol-d4 at 500 MHz and (C) HR-MAS 1H NMR of PHEAA-NH2 microgels in D2O measured with a 4 mm FASTnano probe with a rotor frequency of 4 kHz and 8 kHz at 700 MHz.

added. After 24 h of reaction at 75 °C, the microgel solution became transparent. F


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Figure 3. (A) TEM image of a PHEAA-NH2 microgel sample after 24 h aminolysis of PDEGAC microgels in dioxane and transfer to water. (B) Histogram analysis of the PHEAA-NH2 microgels seen in TEM images, obtained from measuring at least 100 individual microgels, using ImageJ software. (C) LCFM image of PHEAA-NH2 microgels labeled with FITC dye and dispersed in phosphate buffer pH 7.2. (D) Z-average diameter of PHEAA-NH2 colloidal hydrogels in 100 mM phosphate buffer pH7.2 at different temperatures measured by DLS. Data were obtained with a Zetasizer. At each temperature, samples were equilibrated for 10 min prior to each measurement. The viscosity of medium at each temperature was corrected according to viscosity changes for DI water.

One can see that there are no significant changes in dz, consistent with the absence of a VPPT in this temperature range. This is the limit of accessible temperatures on the Zetasizer instrument. To examine higher temperatures, samples of the colloidal PHEAA-NH2 hydrogel solution were sealed in pressure-tested glass vials and heated for detection of a cloud point, a signature of polymer precipitation. Samples at 5 mg/ mL showed no sign of visible turbidity up to 140 °C. These measurements suggest that PHEAA has no LCST up to this temperature in water and that colloidal PHEAA-NH2 hydrogels have sufficient colloidal stability for high-temperature processes. Amine Content of the PHEAA-NH2 Microgels. We used three independent analytical methods to measure the primary amine content of the PHEAA-NH2 microgels. From the HRMAS NMR spectrum of the hydrogels in D2O (see the inset to Figure 2C), we compared the integral of the signals in the range 2.9−3.4 ppm for peaks e, f, and g due to N−CH2 groups adjacent to the amine and amide groups, with the well-resolved broad singlet h associated with the HO−CH2− groups of the hydroxyethylamide pendants. In this way, we found 9 mol % -NH2 in the microgel. By potentiometric and conductometric titration (see Figure S5), in which the microgels were treated with a small excess of HCl and then back-titrated with NaOH, we found 8.6 mol % -NH2 groups. A somewhat smaller value (7.5 mol %), which we consider less reliable, was obtained by a colorimetric assay based on the reaction with picrylsulfonic acid (TNBS assay) (Figure S6). Details of these analyses are discussed in the Supporting Information, and these values are summarized in Table S1.

PHEAA-NH2 microgels accompanied by solvent swelling. The DLS measurements were performed at 25 °C, above the VPTT of PDEGAC particles (15 °C). Thus, the hydrodynamic diameter obtained for PDEGAC in water corresponds to the collapsed state of these particles. DLS measurements were also performed on the microgels in dioxane prior to and after the aminolysis reaction (Table 1). The average diameter for PDEGAC particles in dioxane was dZ = 503 nm, somewhat larger than the diameter of same PDEGAC particles in water at 25 °C (449 nm). The increase in diameter of the PDEGAC particles indicates that dioxane is a better solvent for this polymer compared to water at temperatures above the VPTT of the microgels. After the aminolysis, the diameter measured for PHEAA-NH2 microgels in dioxane was dZ = 913 nm. The chemical transformation of PDEGAC to PHEAA made microgels more swellable in dioxane. To verify the size and structure of the PHEAA-NH2 microgels in water, some of the free amines in the microgel were reacted with FITC. These now fluorescent microgels could be visualized by LCFM, as shown in Figure 3, panel C. The apparent diameter obtained from the LCFM images (ca. 800 nm) is in accord with the diameter obtained by DLS. The histogram analysis of microgels visualized by LCFM is plotted in Figure S4. In addition, in the LCFM image, PHEAA colloidal hydrogels appear to have a fuzzy texture, which is similar to the hairy structure observed by TEM in Figure 3, panel A. The hydrodynamic diameters of PHEAA-NH2 colloidal hydrogels were measured by DLS at various temperatures from 10−70 °C. These values are plotted in Figure 3, panel D. G


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Scheme 2. Chemical Structure of HRP Enzyme Covalently Attached to PHEAA-NH2 Microgels. (A) PHEAA-EDC Microgels. PHEAA-NH2 Microgels Were Treated with Excess Succinic Anhydride To Convert All Amino Groups to Carboxylic Acids. After Activation with EDC, Microgels Were Incubated with HRP. (B) PHEAA-BAH Microgels. PHEAA-NH2 Microgels Were Reacted with S-HyNic. In Parallel, HRP Was Reacted with S-4FB. Modified Microgels and Modified Enzyme Were Mixed and Allowed To React at pH 5.0

Enzyme Immobilization. We used two types of chemistry to attach HRP to the microgels. In the first method, denoted PHEAA-EDC, the amino groups in the PHEAA-NH2 microgels were converted to carboxylic acid functionalities by reaction with excess succinic anhydride. These −COOH groups were in turn activated by reaction with EDC, and after removal of the excess EDC, were reacted with HRP. The product of this synthetic approach is presented in Scheme 2, panel A. For the second immobilization method, we used a bis-aryl-hydrazone (BAH) linkage between the microgel and the enzyme,59 PHEAA-BAH, which is depicted in Scheme 2, panel B. In this chemistry, PHEAA-NH2 microgels were initially modified with S-HyNic. In a separate vessel, the HRP enzyme was reacted with S-4FB. After the unreacted molecules were removed, PHEAA-HyNic and HRP-4FB were incubated together and allowed to react at pH 5.0. As described in the following, this reaction could be monitored by the UV absorption of the product bis-aryl hydrazone. Quantifying Enzyme Loading. We used the Bradford assay to quantify the amount of enzyme immobilized in the microgels. Although this assay is commonly used to evaluate protein content, we had to calibrate the assay to account for a background signal from the microgels. These calibration data are presented in Figure S7 and Table S2, and the results are summarized in Table 2. As shown in Table 2, EDC coupling leads to a slightly higher enzyme loading (35 μg HRP/mg dry microgel) compared to BAH coupling (29 μg HRP/mg dry microgel). Values reported for protein loading in the literature vary substantially, reflecting differences in bead type, protein

dimensions, immobilization chemistries, and reaction conditions. Among the studies in which HRP enzyme was covalently attached to spherical beads, the reported amounts of HRP loading ranged from 7 ng of HRP to 80 μg of HRP per mg of beads (dry weight).64−68 In most cases, high levels of enzyme loading were accompanied by a decrease in enzymatic activity. Porous materials, in principle, should allow even higher levels of protein loading. Recently, Bayne et al.69 reviewed this literature and examined the effect of pore size on the enzyme loading. In this review, the authors assigned different confidence ratings to the various publications, assessing the measurements of porosity and protein loading data for each paper. On the basis of their evaluation, the reliable values of protein loading in porous supports varied between 1 and 500 μg protein per mg support, with the highest loading at about 50 nm pore size. In our microgels, the number of covalently bound HRP enzymes per microgel may be limited by the relatively high cross-link density of microgels caused by the use of 6 wt % BIS as a cross-linker in the synthesis of the precursor particles. It is possible that high cross-linking leads to a small pore size, which limits the penetration of HRP molecules deep inside the microgels. For the PHEAA-BAH sample, we were able to measure the amount of HRP immobilization independently by monitoring the increase in UV absorbance at 354 nm (ε354 = 29 000 M−1 cm−1) during the bioconjugation reaction. The amount of immobilized enzyme in PHEAA-BAH calculated from the UV signal evolution was 22 μg of HRP/(mg microgel) after 4 h of reaction. This value is lower than that obtained by the Bradford assay after sample isolation (29 μg). The difference in these values could be due to either the presence of nonspecifically entrapped HRP in the microgel or the corrections for the microgel contribution to the Bradford assay signal (Table S2). To obtain further information about the enzyme immobilization reaction for PHEAA-BAH, the HyNic-decorated PHEAA microgels were titrated with 4-formylbenzoic acid. The titration results indicated that 31 500 HyNic groups were available per PHEAA microgel. From the bioconjugation value (22 μg HRP/ mg dry microgel), we calculated a mean HRP content of 6200 HRP molecules/microgel particle. Thus, of the 31 500 HyNic groups/microgel particle available as attachment sites, only 20% of these HyNic groups reacted with 4FB-modified HRP. As mentioned previously, by decreasing the cross-link density of our microgel, the pore size should increase, which should enhance the enzyme loading. For comparison, we note that von

Table 2. Amount of HRP Loaded in PHEAA-EDC and PHEAA-BAH Microgels and a Comparison of Kinetic Parameters of Microgel-Bound and Free Enzyme sample

HRP loadinga

Km (μM)

Vmax (μM/min)

kcat (s−1)

native HRP PHEAA-EDC PHEAA-BAH

35 ± 1 29 ± 1

276 290 318

31 27 30

258 225 250

a Bradford assay, μg protein/mg of dry PHEAA. bThe kinetic parameters Vmax, Km, and kcat were obtained from fitting the ABTS assay data with the Michaelis−Menten model. All assays were performed at the same HRP concentration. Each experiment was carried out twice. Calculated values of the parameters for each pair of experiments agreed to better than 5%.

H


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Biomacromolecules Klitzing and co-workers30 entrapped HRP in PNIPAm microgels with a water-swollen diameter of about 1 μm. By using a Bradford assay, they reported a mean value of about 8100 HRP enzyme molecules per microgel particle. In our experiments, we tried to limit the attachment chemistry to one site per enzyme molecule. By titration of the 4FB-modified HRP enzyme with 2-hydrazinopyridine, we could show that about 86% of the HRP molecules have only one 4FB substituent, while the remaining 14% have two or more 4FB substituents. The BAH chemistry guarantees that the vast majority of HRP molecules are attached to the microgel by only one covalent bond. HRP Kinetic Study. Once the amount of enzyme immobilized on the microgels was determined, we studied the kinetics of the immobilized HRP enzyme and compared its catalytic activity to that of the native enzyme. The substrates used for this study were H2O2 and 2,2′-azino-bis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS), and measurements were performed in 50 mM phosphate buffer pH 5.2. Initial rates were measured, and the kinetic parameters, Km, Vmax, and kcat, were obtained from fitting the kinetic data to the Michaelis− Menten kinetic model (Figure S8). The results from these studies are summarized in Table 2. From these data, we see that the Km values for the PHEAA-BAH and PHEAA-EDC samples are slightly higher than that of the native enzyme, suggesting that the microgel-bound enzyme has a lower affinity toward its substrates. A lower local concentration of the substrates in the microgel and limited mass transfer within the microgel boundaries might cause this increase in Km. Nevertheless, both bound enzyme and the free enzyme have similar values of kcat, demonstrating that HRP has maintained its activity upon immobilization. HRP Thermal Stability. The goal of this research was to synthesize aqueous microgels that show no thermal transitions in water and to examine whether these aqueous microgels are useful as a support for high-temperature enzymatic reactions. HRP is known to have its maximum activity above room temperature. Thus, it provides a useful test for our microgels. We chose the PHEAA-BAH sample for these tests and measured the activity of native HRP and immobilized HRP at different temperatures in the range of 10−70 °C. The results of these experiments are shown in Figure 4. In our experiments, the maximum activity measured in phosphate buffer pH 5.2 for both native and immobilized HRP was about 40 °C. At most temperatures, the activity of native HRP was very similar to that of PHEAA-HRP, but at the highest temperatures (i.e., 60, 70 °C), the activity of PHEAA-HRP was higher than that of native HRP. The similarity in the trends and the values of enzyme activity indicate that the microgel as an enzyme support did not interfere with the enzyme performance. The colloidal stability and pore size of PHEAA microgels, which control the rate of mass transfer, appear to have remained unchanged at all of these temperatures. The lower activity observed for the native HRP at 60 and 70 °C is likely due to denaturation of the native enzyme upon heating to those temperatures. Our results stand in strong contrast to experiments in which enzymes were attached to stimuli-responsive polymers and examined at elevated temperatures. In terms of enzymes that prefer a hydrophilic environment, heating the solution of immobilized proteins on a thermoresponsive support beyond the LCST of their support led to a decrease in bioactivity. For example, Hoffman and coauthors, in a series of publications,70−73 prepared various stimuli-responsive polymer−

Figure 4. Activity of native HRP (blue empty squares) and immobilized HRP (PHEAA-HRP) (red filled diamonds) at different temperatures. The ABTS assay was performed at the same HRP concentration for both samples. For each assay, after the sample reached thermal equilibrium at each temperature, H2O2 was injected, and the change in absorbance at 405 nm was recorded. The rate for each enzyme at each temperature was measured twice. All values of the activity for each pair of measurements agreed to better than 2%.

enzyme conjugates in which the enzymatic activity was totally “switched-off” after the polymer collapse. Russell and coworkers74,75 used a “growing-from” approach to attach multiple copies (10−14) of thermoresponsive (or pH responsive) polymers to chymotrypsin as a prototypical enzyme with the idea of using the thermal or pH response to modulate enzyme activity. They found, for example, that above the LCST of their polymers, Km increased for the hydrophobic peptide substrates, but catalytic efficiency, as measured by the ratio kcat/Km, decreased compared to the native enzyme. The thermal stabilities of the native enzyme and the microgel-conjugated enzyme were examined at 50 °C, and the results are presented in Figure 5. Both the native HRP and immobilized HRP (PHEAA-BAH sample) were annealed at 50 °C in phosphate buffer at pH 6.8. Samples were withdrawn at

Figure 5. Residual activity of native HRP (red filled diamonds) and immobilized HRP (PHEAA-HRP) (blue empty squares) after annealing of the enzyme at 50 °C in phosphate buffer pH 6.8. The activity measurement was performed at 25 °C by recording the change in absorbance of ABTS assay at 405 nm. For each sample, the obtained activity was normalized to the maximum value (time = 0). I



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30 min intervals, and their kinetics were measured at room temperature. The results showed a substantial difference in the residual activities of native and immobilized HRP. After 330 min of annealing at 50 °C, the microgel-bound enzyme recovered 76% of its initial activity, whereas native HRP recovered only 20% of its original activity. The enzyme activity of PHEAA-HRP showed a more pronounced decrease over the first 30 min and then exhibited a more linear degradation rate. In contrast, native HRP demonstrated a very sharp decay of activity over time. The decay rates of native and immobilized HRP activity were fitted to an exponential profile using eq 2, by assigning t0 to the data points at 30 min of exposure to 50 °C (Figure S9). It is not meaningful to include the data points at zero annealing time in these fits because these rates refer to enzyme solutions that were not treated at 50 °C. In this way, we obtained decay rates at 50 °C of kdecay‑HRP = 4.2 × 10−3 min−1 for the free enzyme and kdecay‑PHEAA‑HRP = 6.5 × 10−4 min−1 for the microgelimmobilized HRP. We find that the decay rate of native HRP is about 6.5-times faster than that of the PHEAA-HRP immobilized enzyme. The data in Figures 4 and 5 show that immobilization of HRP on a microgel support increased its stability at high temperatures compared to the native HRP. We interpret this result to suggest that covalent bonding between HRP and the microgel suppressed enzyme aggregation and increased the enzyme resistance against unfolding during the annealing.

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CONCLUSIONS

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ASSOCIATED CONTENT

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. 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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We report the design and synthesis of a functional aqueous microgel sample that showed no changes in volume upon heating and can be used as a support for biomacromolecule attachment. We used a two-step process to synthesize these PHEAA-NH2 microgels by starting with the preparation of PDEGAC microgels in water, transfer to dioxane, and then exhaustive aminolysis of the acrylate ester groups with ethanolamine plus a small amount of ethylenediamine. The PHEAA-NH2 microgels maintained their hydrodynamic diameter up to 70 °C, the maximum temperature accessible with our Zetasizer, and exhibited no cloud point up to 140 °C. By taking advantage of the amine functionality of the microgels, two different chemistries were used to covalently bind HRP as a model enzyme. The PHEAA microgels proved to be efficient at stabilizing the enzyme at high temperatures (up to 70 °C) without retarding the enzymatic activity. These thermally stable microgels show great promise as supports for thermophilic enzymes and for the support of other biomacromolecules employed in high-temperature reactions in water.

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.biomac.5b00768. Experimental details and protocols, SEM images, 1H NMR spectra, DLS CONTIN plots and autocorrelation decay profiles, titration curves, TNBS assay data, Bradford assay data, enzyme kinetic plots, enzymatic activity decay profiles (PDF) J


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