Polymeric Nanocapsules for Enzyme Stabilization in Organic Solvents

Jan 2, 2018 - (18-21) However, the method still requires the chemical modification of the enzyme surface with polymerizable vinyl groups to create a p...
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Article Cite This: Macromolecules XXXX, XXX, XXX−XXX

Polymeric Nanocapsules for Enzyme Stabilization in Organic Solvents Fumi Ishizuka,† Robert Chapman,‡,∥ Rhiannon P. Kuchel,§ Marion Coureault,‡ Per B. Zetterlund,*,† and Martina H. Stenzel*,‡ †

Centre for Advanced Macromolecular Design, School of Chemical Engineering, ‡Centre for Advanced Macromolecular Design, School of Chemistry, ∥Australian Centre for Nanomedicine, School of Chemistry, and §Electron Microscope Unit, Mark Wainwright Analytical Centre, The University of New South Wales, Sydney, NSW 2052, Australia S Supporting Information *

ABSTRACT: Herein we report an approach to encapsulate enzymes within polymeric nanocapsules dispersed in an organic solvent via inverse miniemulsion periphery RAFT polymerization (IMEPP). Glucose oxidase (GOx), which has various applications but is unstable at elevated temperature and in organic solvents, was chosen as a model enzyme. In this study, we have explored the use of photoinitiation under visible (blue) light instead of thermal initiation to avoid enzyme denaturation by heating. GOx was successfully encapsulated within polymeric nanocapsules (∼200 nm) and showed high activity (71−100% relative to free GOx in PBS) dispersed in toluene/t-BuOH. The nanocapsules were thus able to protect GOx and enable it to function in an organic solvent mixture where native GOx would otherwise undergo denaturation. This approach of enzyme encapsulation is significant as it may lead to increased industrial applications of enzyme biocatalysis, expanding the use of enzymes as nontoxic and environmentally friendly biocompatible catalysts.

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INTRODUCTION Enzymes have been utilized as biocatalysts in a wide range of industries such as the food and pharmaceutical industries due to their ability to catalyze various reactions at high specificity under mild conditions as well as their biocompatibility and biodegradability.1−4 However, industrial use of enzymes is still challenging due to the intolerance of enzymes to harsh conditions such as high temperatures and organic solvents. Enzymes are soluble and active under aqueous conditions, but water is not a suitable solvent to solubilize various substances such as pharmaceutical intermediates. Although enzymes are usually insoluble and inactive in organic solvents,5 the use of enzymes in organic solvents offers various advantages such as improved solubility of hydrophobic substrates and minimization of side reactions that occur in water. As such, a number of researchers in both industry and academia have been trying to stabilize enzymes in organic solvents.6,7 Various strategies have been reported such as chemical modification, absorption, or entrapment onto/in an inert matrix and reverse micelles. Chemical modification of enzymes is a popular way to stabilize enzymes in nonaqueous environments.7 The surface modification of enzymes with polymers such as poly(ethylene glycol) (PEG) has been widely studied.8,9 Covalently crosslinked enzymes such as cross-linked enzyme aggregates (CLEA) have also demonstrated increased tolerance to organic solvents.10 However, such chemical modification of enzymes is often associated with an increased risk of altering the original conformation, which could decrease the enzymatic activity. © XXXX American Chemical Society

Physical entrapment of enzymes in reverse micelles/microemulsions is another common approach used to stabilize enzymes in organic solvents;11−13 however, this technique is associated with the risk of enzyme inactivation by surfactants.7 The physical encapsulation of enzymes in polymer matrices has a range of advantages such as easy separation of products by filtration or centrifugation while the collected enzymes can be reused. Enzyme-loaded nanocapsules have been extensively studied for protein delivery for biomedical applicationsvarious techniques to achieve enzyme stabilization via encapsulation within polymeric nanocapsules have been reported, including the use of polymer micelles, liposomes, and nanoparticles.14−16 These systems were designed for dispersions in physiological environments, i.e., in water, and are therefore not suitable for enzyme stabilization in nonaqueous systems. Despite recent developments in the field of nanocapsule fabrication for protein delivery, the preparation of enzyme-loaded polymeric nanocapsules is still challenging due to poor stability of enzymes.14−16 Yang et al. reported successful stabilization of enzymes by radical polymerization from the enzyme surface, resulting in good enzyme activity in a water/organic solvent mixture.17 This approach, referred to as “single-enzyme nanogel”, has been applied for various enzymes such as Received: November 7, 2017 Revised: December 9, 2017

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DOI: 10.1021/acs.macromol.7b02377 Macromolecules XXXX, XXX, XXX−XXX

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Scheme 1. Schematic Illustration of Synthesis of Nanocapsules via Inverse Miniemulsion Periphery RAFT Polymerizationa

a

The dispersed phase (blue) contains the lipophobe NaCl and the enzyme GOx, and the continuous phase (gray) initially contains amphiphilic macroRAFT agent as colloidal stabilizer as well as mono- and divinyl monomer.

catalase.18−21 However, the method still requires the chemical modification of the enzyme surface with polymerizable vinyl groups to create a polymeric shell around the enzyme. Such chemical modification of enzymes can result in unpredictable risks of alternation of the original conformation of enzymes, leading to reduced enzymatic activity. In addition, such chemical modifications usually utilize amino acid residues on the protein surface. These functional groups can be different from enzyme to enzyme, requiring optimization steps for each enzyme. Physical encapsulation in nanocapsules without chemical modification is a more versatile way to stabilize enzymes regardless of their structure. The polymeric shells act to prevent enzymes from leaching out while allowing small molecule substrates to penetrate into the core. Because of the high surface area, substrates can diffuse into the core at sufficient rates where enzymes are entrapped, enabling them to function as nanobiocatalysts.22−24 However, there is a need to develop methodology that is applicable to organic solvents. We have previously reported a synthetic pathway to polymeric nanocapsules with a hydrophilic core via inverse miniemulsion periphery RAFT polymerization (IMEPP).25−30 The method is based on self-assembly of an amphiphilic macroRAFT agent at the water droplet interface, followed by RAFT polymerization of mono- and divinyl monomer located in the hydrophobic continuous phase leading to formation of a thin polymer shell around the water droplet. The success of this pathway is based on the availability of the RAFT end groups and the high end group fidelity of the RAFT polymer.31 Successful encapsulation and release of a model protein (BSA)27 and hydrophilic anticancer drug (gemcitabine)26 have been demonstrated using this technique. The IMEPP approach has great potential for protein encapsulation as the polymerization reaction occurs at the periphery of the water droplets as opposed to within the droplets where the guest molecules are located. However, to date, the IMEPP process has been carried out at 60−70 °C relying on thermal initiation, and as such the process is limited to the encapsulation of robust molecules. To encapsulate temperature sensitive molecules such as enzymes without denaturation occurring, a milder approach is required. Herein, we report the synthesis of nanocapsules via IMEPP under visible light (Scheme 1), demonstrating successful encapsulation of a model enzyme which is unstable at elevated temperatures. Enzyme-loaded nanocapsules were synthesized via IMEPP using the photoinitiation system camphorquinone (CQ) and ethyl 4-(dimethylamino)benzoate (EDB) under visible light (ultraviolet light, which is often used for photopolymerization, can deactivate enzymes32,33). Glucose oxidase (GOx) was chosen as a model enzyme to be

encapsulated as its activity can easily be monitored using a colorimetric assay, while it is also a widely used enzyme that is easily denatured at higher temperatures.

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EXPERIMENTAL SECTION

Materials. All materials were reagent grade and used as received, unless otherwise specified: Di(ethylene glycol) methyl ether methacrylate (DEGMA, 95%, Sigma-Aldrich), sodium chloride (NaCl, Univar), camphorquinone (CQ, Sigma-Aldrich), ethyl 4(dimethylamino)benzoate (EDB, >98.0%, TCI), Nile Blue A (NB, Sigma-Aldrich), glucose oxidase from Aspergillus niger (GOx, SigmaAldrich), bovine serum albumin (BSA, Sigma-Aldrich), horseradish peroxidase (HRP, Sigma-Aldrich), 3,3′,5,5′-tetramethylbenzidine (TMB, Sigma-Aldrich), sulfuric acid (H2SO4, Sigma-Aldrich), Dglucose (Sigma-Aldrich), diethyl ether (Et2O, 99%, Univar), toluene (99.5%, Univar), methanol (Chem supply), n-hexane (Chem supply), tert-butanol (t-BuOH, Ajax Funechem), tetrahydrofuran (THF, Sigma-Aldrich) and N,N-dimethylacetamide (DMAc, 99.9%, SigmaAldrich) were used without further purification. Deuterated NMR solvents (CDCl 3) were purchased from Cambridge Isotope Laboratories. Methyl methacrylate (MMA, >99%, Sigma-Aldrich), lauryl methacrylate (LMA, 96%, Sigma-Aldrich), tert-butyl methacrylate (tBMA, 98%, Sigma-Aldrich), and ethylene glycol dimethacrylate (EGDMA, 98%, Sigma-Aldrich) were deinhibited by passing through a column of activated basic alumina. Deinhibited monomers were stored below 4 °C. 2,2′-Azobis(isobutyronitrile) (AIBN) was recrystallized twice from methanol. Deionized (DI) water was produced by a Milli-Q reverse osmosis system and had a resistivity of 19.6 mΩ cm−1. The RAFT agent, 4-cyanopentanoic acid dithiobenzoate (CPADB), was synthesized according to the literature.34 Analyses. Size Exclusion Chromatography (SEC). SEC was performed using a Shimadzu modular system, comprising an SIL10AD autoinjector, an LC-10AT pump, a DGU-12A degasser, a CTO-10A column oven, and an RID-10A differential refractive index detector. A column arrangement consisting of a Polymer Laboratories 5.0 μm bead size guard column (50 × 7.8 mm) followed by four linear PL column (300 × 7.8 mm, 500, 103, 104, and 105 Å, 5 μm pore size) was used for the analysis. DMAc was used as the mobile phase at a constant temperature of 50 °C and a constant flow rate of 1 mL min−1. The SEC system was calibrated using linear poly(methyl methacrylate) standards, ranging from 500 to 106 g mol−1 (Polymer Laboratories). THF was used as the mobile phase to analyze pLMA based polymers at 40 °C flow rate of 1 mL min−1. The THF system was calibrated against linear poly(styrene) standards ranging from 500 to 105 g mol−1 (Polymer Laboratories). Chromatograms were processed using Cirrus 2.0 software (Polymer Laboratories). Nuclear Magnetic Resonance (NMR). NMR was utilized to analyze the structure of the synthesized compounds as well as to determine the monomer conversion in the block copolymer synthesis and nanocapsule synthesis. 1H NMR spectroscopy was carried out using a Bruker Avance III HD 400 MHz, equipped with an autosampler system. Chemical shifts are reported in parts per million B

DOI: 10.1021/acs.macromol.7b02377 Macromolecules XXXX, XXX, XXX−XXX

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Synthesis of PDEGMA-b-PLMA MacroRAFT Agent. Poly(DEGMA) macroRAFT agent (1.38 g, 2.63 × 10−4 mol), LMA (13.39 g, 5.26 × 10−2 mol), and AIBN (0.0043 g, 2.63 × 10−5 mol) were dissolved in 1,4-dioxane (15 mL) to give [monomer]:[RAFT]: [initiator] = 200:1:0.1. The solution was thoroughly degassed in an ice bath for 30 min before being placed in an oil bath at 70 °C for 6 h. The polymerization was stopped by placing the solution in an ice bath for 30 min. The final solution was then precipitated in methanol to yield a pink viscous liquid. The monomer conversion was 29% by 1H NMR (Mn,th = 18 600 g mol−1, Mn,sec = 22 900 g mol−1 (with respect to PMMA standards), Đ = 1.15). 1H NMR (400 MHz, CDCl3): δ (ppm) = 3.8−4.0 (2nH, −COOCH2C11H23), 2.1−1.2 (2nH, CH2 of the main chain and −COOCH2C10H20CH3), 1.1−0.7 (3nH, CH3 of main chain and −COOCH2C10H20CH3). Preparation of Inverse Miniemulsions. Inverse miniemulsions were prepared using the recipe listed in Table 2. A representative emulsion preparation process is as follows: the macroRAFT was dissolved in toluene (8.67 g) to create the continuous phase of the emulsion. In a separate vial, distilled water (0.434 mL, 5 wt % relative to toluene) and NaCl (0.0173 g, 4 wt % relative to water) were mixed to create the dispersed phase and then poured into the vessel. To check the permeability of the nanocapsules shells, Nile Blue A was incorporated in nanocapsules as a guest molecule. For encapsulation of Nile Blue, Nile Blue was dissolved (1.6 mg/mL) in the dispersed phase (2 wt % NaCl aqueous solution). For enzyme encapsulation, GOx was dissolved in the dispersed phase (PBS) as a model enzyme (1 mg/mL in PBS). BSA (5 mg/mL) was added to the dispersed phase to prevent absorption to surfaces such as glass and the sonicator tip. Both phases were mixed and subsequently homogenized by ultrasonication (Branson 450 sonifier, 55% amplitude, 5 min, 5 mm tip diameter). Inverse Miniemulsion Periphery RAFT Polymerization. For a typical cross-linking reaction via inverse emulsion periphery RAFT polymerization, a solution consisting of t-BMA (monomer, M; 0.1006 g; 0.71 × 10−3 mol), EGDMA (cross-linker, XL; 0.0175 g, 0.88 × 10−4 mol), CQ (initiator, I1; 0.6 mg, 3.54 × 10−6 mol), and EDB (coinitiator, I2; 2.3 mg, 1.2 × 10−4 mol) was added to the mixture after emulsification to give a [M]:[XL]:[RAFT]:[I1]:[I2] ratio of 200:25:1:1:3.4 (Table 3). The resulting mixtures, prepared in accordance with the procedure described above, were degassed by purging with nitrogen in an ice bath for 30 min. Polymerization was carried out at room temperature for 1 h under blue LED light (intensity 45 mW/cm2) with constant stirring. The emission spectrum of the blue LED is shown in Figure S1. After polymerization, 1 mL of the emulsion was purified according to the following (removing unreacted monomer and initiator and dried under vacuum.): R1 and R2: precipitation in excess MeOH followed by centrifugation (7000 rpm for 5 min) or centrifugation (14 500 rpm for 30 min) without addition of MeOH; R3 and R4: precipitation in excess hexane or MeOH (both approaches used for both R3 and R4) followed by centrifugation (7000 rpm, 5 min). The dried samples were redispersed in 1 mL of toluene by shaking (no ultrasonication). DLS and TEM analyses were carried out on the diluted emulsion (0.1 mL of emulsion in 2 mL of solvent). Release Study of Nile Blue from Nanocapsules. The release of NB from nanocapsules was studied by fluorescence spectroscopy (Cary Eclipse) to investigate the permeability of the polymeric shell for small molecules. The NB-loaded nanocapsules were diluted with toluene and t-BuOH to mimic the condition for enzyme assay then centrifuged 14 500 rpm for 30 min, and the fluorescence spectra of NB from supernatant and the NB-loaded nanocapsules (without centrifuge) were measured. Measurement of GOx Activity. The activity of encapsulated GOx was measured by colorimetric assay relative to a standard curve of GOx in PBS. Samples of dispersed nanocapsules in toluene at varying dilutions (75 μL), or of a standard curve of GOx in PBS (0−5 μg/mL, 75 μL), were incubated at 30 °C for 15 min after addition of 25 μL of glucose (100 mM in t-BuOH). The amount of H2O2 generated by the GOx over this time was then measured using an excess of horseradish peroxidase (HRP). This was done by diluting

(ppm), relative to the residual solvent peak. The theoretical molecular weight (Mn,th) was calculated according to eq 1:

M n,th =

[monomer] × conversion × M monomer + MRAFT [RAFT]

(1)

where MRAFT denotes the molecular weight of the RAFT agent or the macroRAFT agent. Particle Size Measurement. DLS measurements were conducted on a Zetasizer Nano ZS (Malvern), with a 4 mV He−Ne laser operating at λ = 632 nm and noninvasive backscatter detection at 173°. Measurements were conducted in a quartz cuvette at 25 °C, with 30 s equilibration period prior to each set of measurements. For a given sample, a total of three measurements were conducted. In each measurement, the number of runs, attenuator, and path length used were automatically adjusted by the instrument, depending on the quality of the sample. The presented results are averages of the three measurements. Transmission Electron Microscopy (TEM). TEM images were obtained by using a JEOL1400 TEM operating at an accelerating voltage of 100 kV. Images were recorded via the Gatan CCD imaging software. All TEM samples were prepared by dropping a 1 mg mL−1 emulsion on a Formvar-supported copper grid. Excess solvent was drained using filter paper after 1 min. Cryo-Transmission Electron Microscopy (Cryo-TEM). Vitrified specimens were analyzed with cryo-TEM using the following method; lacey carbon grids (Pro Sci Tech, QLD, Australia) were hydrophilized by glow discharge for 60 s (Pelco easiglow; Ted Pella Inc., Redding, CA). Specimens were then prepared within a controlled environment using the Leica EM GP (Leica, NSW, Australia), relative humidity 99% at 25 °C. Two microliters of the sample solution was then pipetted onto the hydrophilized lacey grid and blotted for 1.5 s. The grid was then automatically plunged into liquid ethane at its freezing temperature (−183 °C) to form a vitrified layer. The vitrified samples were examined using a FEI Tecani (FEI, OR, USA), operating at an accelerating voltage of 200 kV. A Gatan 626 cryo holder (Gatan, Pleasanton, CA), was used to maintain the sample temperature below −172 °C during both the transfer and imaging processes. Images were recorded digitally using an Eagle 2k CCD camera (FEI) and Digital Micrograph (Gatan). All samples were analyzed using the low-dose software (FEI) to minimize beam exposure and electron beam radiation damage. Synthesis of PDEGMA MacroRAFT Agent. DEGMA (10 g, 5.369 × 10−2 mol), CPADB (0.3 g, 1.074 × 10−3 mol) as RAFT agent, and AIBN (0.0176 g, 1.074 × 10−4 mol) as initiator were dissolved in toluene (21.5 mL) to give a [monomer]:[RAFT]: [initiator] molar ratio of 50:1:0.1. The reaction mixture was thoroughly purged with nitrogen gas for 30 min before being placed in an oil bath at 70 °C for 4 h. After polymerization, the reaction was stopped by placing the mixture in an ice bath for 30 min. The polymer was isolated by precipitation in n-hexane to yield poly(DEGMA) as a viscous red liquid. The monomer conversion was determined to be 35% via 1H NMR (Mn,th = 3500 g mol−1, Mn,SEC = 4400 g mol−1 (with respect to PMMA standards), Đ = 1.1). 1H NMR (400 MHz, CDCl3): δ (ppm) = 4.1 (2nH, −COOCH2−), 3.7−3.5 (6nH, −CH2O), 3.4 (3nH, −OCH3), 2.1−1.7 (2nH, CH2 of the main chain), 1.1−0.9 (3nH CH3 of the main chain), where n is the degree of polymerization. Synthesis of PDEGMA-b-PMMA MacroRAFT Agent. Poly(DEGMA) macroRAFT agent (1.23 g, 3.47 × 10−4 mol), MMA (6.96 g, 6.95 × 10−2 mol), and AIBN (0.0057 g, 3.47 × 10−5 mol) were dissolved in toluene (7 mL) to give [monomer]:[RAFT]:[initiator] = 200:1:0.1. The solution was thoroughly degassed in an ice bath for 30 min before being placed in an oil bath at 60 °C for 18 h. The polymerization was stopped by placing the solution in an ice bath for 30 min. The final solution was then precipitated in diethyl ether to yield a brittle, pink solid. The monomer conversion was 44% by 1H NMR (Mn,th = 12 300 g mol−1, Mn,sec = 16 000 g mol−1 (with respect to PMMA standards), Đ = 1.04). 1H NMR (400 MHz, CDCl3): δ (ppm) = 3.6 (3nH, −COOCH3), 1.9−1.8 (2nH, CH2 of the main chain), 1−0.8 (3nH, CH3 of main chain). C

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18 600) and 16 000 (M n,th = 12 300) g mol −1 and polydispersities (Đ) of 1.15 and 1.04, respectively. The HLB values were 5.8 (macroRAFT1) and 5.2 (macroRAFT2), respectively, calculated as described previously.30 HLB values of steric stabilizers for inverse miniemulsions are usually in the range 3−8.35 Preparation of Inverse Miniemulsions. Inverse miniemulsions were prepared by ultrasonication using macroRAFT1 and macroRAFT2 as a steric stabilizers (Table 2).

the samples 5 times in t-BuOH and mixing 10 μL of this diluted solution with 90 μL of HRP (5 μg/mL) and 3,3′,5,5′-tetramethylbenzidine (TMB, 1 mM) in citrate buffer (50 mM, pH 5.5). The reaction was quenched after 1 min with H2SO4 (20 μL, 1 M), and the absorbance at 450 nm was measured. The activity of GOx in the particles was determined using eq 2:

GOx activity (%) =

[GOx]active [GOx]initial

(2)

where [GOx]initial is the theoretical concentration of GOx in the miniemulsion based on the amount of GOx been added before crosslinking and [GOx]active is the concentration of active GOx measured in the activity assay, as calculated from the eq 3 based on the standard curve of GOx in PBS (Figure S4). [GOx]active = (Abs − 0.0462)/0.0434

Table 2. Inverse Miniemulsion Conditions Used To Synthesize GOx-Loaded Nanocapsules cont phase

(3)

disp phase

toluene (g) macroRAFT (g)

To study the effect of ultrasonication and light irradiation on GOx activity, GOx solution (1 mg/mL in PBS with 5 mg/mL BSA) was exposed to ultrasonication or light irradiation (UV, blue LED) in the same way as during the nanocapsule preparation.

DI water (mL) NaCl (mg) PBS (mL) GOx (mg) BSA (mg) ultrasonication (time, amplitude, power)

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RESULTS AND DISCUSSION Synthesis of MacroRAFT Stabilizers. Amphiphilic diblock copolymers with a suitable hydrophilic−lipophilic balance (HLB) were synthesized via RAFT polymerization. Two types of macroRAFT agents, poly(di(ethylene glycol) methyl ether methacrylate)24-b-poly(lauryl methacrylate)54 (“macroRAFT1”) and poly(di(ethylene glycol) methyl ether methacrylate) 17 -b-poly(methyl methacrylate)87 (“macroRAFT2”), were synthesized (number of repeating units from 1 H NMR). Di(ethylene glycol) methyl ether methacrylate was chosen as hydrophilic segment because of its good solubility in the continuous phase (toluene) as well as its hydrophilicity, while LMA and MMA were selected as hydrophobic segments with low and high Tg, respectively, to investigate the effect of the polymer type on the permeability of the polymeric shell of the nanocapsules. SEC analyses of macroRAFT1 and macroRAFT2 (Figure 1 and Table 1) gave Mn = 22 900 (Mn,th =

without GOx

with GOx

8.67 0.087 0.434

8.67 0.087

0.0173

comment 1 wt % rel to toluene 5 wt % rel to toluene 4 wt % rel to water

0.434 0.4

5 wt % water content rel to toluene 1 mg/mL in PBS

2.2

5 mg/mL in PBS 5 min, 55%, 450 W

GOx was dissolved in the dispersed phase (phosphate buffered saline, PBS) before encapsulation by ultrasonication with the macroRAFT agent in toluene. The intensity size distributions of the initial droplets (by dynamic light scattering, DLS) for both macroRAFT agents with/without GOx are displayed in Figure 2kinetically stable miniemulsions were obtained in all

Figure 1. Molecular weight distributions (“GPC distributions”, i.e., w(log M) on y-axis, normalized to peak height) of macroRAFT1 (full line) and macroRAFT2 (dotted line).

Figure 2. Normalized intensity based size distributions of inverse miniemulsions (before polymerization) stabilized by macroRAFT1/2 with/without GOx.

Table 1. Synthesis Parameters and Molecular Weight Data of Amphiphilic MacroRAFT Agents P(DEGMA24-b-LMA54) “macroRAFT1” P(DEGMA17-b-MMA87) “macroRAFT2”

[M]/[RAFT]/[AIBN]

time (h)

conv (%)

Mn,th (g mol−1)

Mn,sec (g mol−1)

Đ

HLBa

200:1:0.1 200:1:0.1

6 18

29 44

18 600 12 300

22 900 16 000

1.15 1.04

5.2 5.8

a HLB values were obtained from the number-average molecular weight ratio of the hydrophilic segment (Mh) and the total molecular weight (M): HLB = 20(Mh/M).

D

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Table 3. Conditions for Synthesis of Nanocapsules via Inverse Miniemulsion Periphery RAFT Polymerization under Visible Light (Blue LED, λmax = 463 nm) with CQ (I1)/EDB (I2) as Photoinitiation System

R1 R2 R3 R4

stabilizer

shell

macroRAFT1d

LMA/EGDMA

macroRAFT2e

t-BMA/EGDMA

[macroRAFT]:[M]a:[XL]b: [I1]:[I2]

time (h)

conv (%)

DH (nm)f

PDIf

shell thicknessc(nm)

[GOx]in PBS (mg/mL)

1:300:100:5:17 1:600:200:5:17 1:200:25:1:3.4 1:600:200:1:3.4

1 1 1 1

26 30 27 24

122 129 322 232

0.093 0.043 0.26 0.11

6.0 12.9 3.5 10.7

1

a

[M] monomer (LMA for R1, R2 and t-BMA for R3, R4). b[XL] cross-linker (EGDMA). cEstimated shell thickness based on volume of added monomer and cross-linker (see Supporting Information). dMacroRAFT1 = P(DEGMA24-b-LMA54) (Mn,SEC = 22 900 g mol−1, Đ = 1.15). e MacroRAFT2 = P(DEGMA17-b-MMA87) (Mn,SEC = 16 000 g mol−1, Đ = 1.04). fIntensity-averaged diameter of nanocapsules in toluene purified by centrifuge (R1 and R2) or precipitation in hexane (R3 and R4) and PDI determined by DLS

termination involving propagating radicals attached to two different particles as well as propagation involving a propagating radical attached to one particle with a pendant vinyl bond of another particle would lead to interparticle crosslinking (aggregation). Both the absolute amounts of monomer and cross-linker as well as the monomer/cross-linker ratio were carefully adjusted, and the overall conversions were deliberately maintained relatively low to minimize such interparticle cross-linking.30 For R2 and R4, higher initial monomer and cross-linker loadings were used than for R1 and R3 to make the polymeric shell thicker. Approximate estimates of the shell thicknesses based on the volume of monomer and cross-linker reacted (calculation outlined in the Supporting Information) are listed in Table 3. In all four runs (R1−R4), the cross-linked capsules exhibited similar size distributions to the initial droplets (Figure 4). After purification (removal of unreacted monomer) by precipitation in a nonsolvent or centrifugation (see Experimental Section for detailed procedures), the nanocapsules were redispersed in toluenealthough the PDI increased somewhat for R3 (approximately 0.3) compared to the others, similar size distributions were confirmed in all cases. The results demonstrate that the nanocapsules were sufficiently cross-linked to maintain their integrity even after purification. Capsule formation was confirmed by TEM imaging. Figure 5 and Figure S2 show GOx-loaded nanocapsules synthesized using the recipes in Tables 2 and 3. In all cases, nanoparticles with hollow structure were observed with diameters in the range 50−250 nm. The nanocapsules appear to be somewhat deflated with relatively thin shells. Because of the insolubility of GOx in the toluene phase, it is believed that GOx was entrapped inside the aqueous core of the nanocapsules, although GOx is difficult to observe by TEM due to its small size (overall dimensions of 6.0 × 5.2 × 7.7 nm3) and low electron density.37 Based on the amount of GOx added and the size of the initial droplet diameters, each nanocapsule is estimated to contain 8−21 GOx molecules, depending on the initial droplet diameter for R1−R4 (120−170 nm by DLS). Permeability of Nanocapsules. Encapsulated enzymes can only provide useful function if the substrate, here glucose, is capable of diffusing through the protective polymer layer. Given that glucose is insoluble in toluene and considering that toluene is of course immiscible with water, a third solvent (tBuOH) was needed in order to transport glucose across the membrane of the nanocapsules into the interior. We were therefore interested in studying how the addition of solvents facilitated transport of cargo across the nanocapsule membrane. For this purpose, Nile Blue (Figure 6D), the fluorescence maximum of which is a function of the polarity of the solvent, was added to the aqueous phase prior to the

cases. The initial droplet diameters were approximately 120− 170 nm with monomodal distributions with low polydispersity index (PDI < 0.1 for all samples). The presence of GOx does not seem to affect the stability of the macroRAFT1/ macroRAFT2 miniemulsions. These results clearly demonstrate the successful containment of GOx in the dispersed phaseGOx is insoluble in toluene and would as such be detected as a secondary peak (precipitate) if present in the continuous phase and no such large-scale precipitate was observed. Synthesis of Nanocapsules via Inverse Miniemulsion Periphery RAFT Polymerization. In our previous work,25−30 IMEPP was conducted at elevated temperatures (60−70 °C) with AIBN as initiator. In the present study, the photoinitiation system CQ/EDB was utilized to carry out IMEPP under light irradiation at room temperature to avoid enzyme denaturation.33 Blue light (λmax = 463 nm) was selected, considering that UV light can deactivate GOx over time (Figure S3B). Two different systems were investigated, namely, LMA/EGDMA (R1 and R2) and t-BMA/EGDMA (R3 and R4) based on the recipes in Tables 2 and 3, designed such that the monomer to be polymerized matches that of the hydrophobic segment of the macroRAFT in terms of the Tg values of the corresponding linear polymers (Tg = −50 (PLMA homopolymer) and 107 °C (PtBMA homopolymer)).36 Polymerizations were first carried out without GOx. Successful nanocapsule formation was confirmed by TEM (Figure 3).

Figure 3. CryoTEM (A) and normal TEM (B) micrographs of polymeric hollow nanocapsules synthesized using the same recipe as R1 (A) and R3 (B) but without GOx. Black objects are NaCl crystals.

Polymerizations were subsequently performed with GOx present in the aqueous phase based on the recipes in Table 3. In all cases, the polymerizations proceeded with good colloidal stability, leading to overall conversions of 26, 30, 27, and 24% for R1, R2, R3, and R4, respectively. Given that these polymerizations occur in the continuous phase, bimolecular E

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Figure 4. Intensity based size distributions (normalized to peak height) of initial droplets, cross-linked capsules (crude), and capsules purified by centrifugation (R1 and R2, Table 3) or precipitation in n-hexane (R3 and R4, Table 3) and then redispersed in toluene.

Figure 6. (A) Photographs of Nile Blue (NB) in water (left), NBloaded nanocapsules (NC) prepared from the raw miniemulsion after polymerization by dilution with t-BuOH to give toluene/t-BuOH = 75/25 vol % (middle), and NC in toluene prepared from the raw miniemulsion after polymerization by dilution with toluene to the same NC concentration as in (B) (right); NB shows blue color in aqueous solution while it turns light pink in toluene/t-BuOH. The concentration of water in the middle and right sample is 0.39 vol %. (B) Fluorescence spectra (Ex 635 nm, Em 640−900 nm) of free NB in various solvents, NB released from NC and NB encapsulated in NC. The concentration of NB is 1.76 × 10−5 mol/L in all samples in A and B (except NB leached from NC). (C) Photograph of NBloaded NC (no dilution). (D) Chemical structure of Nile Blue.

Figure 5. TEM micrographs of GOx-loaded polymeric nanocapsules from reaction (A) R1, (B) R2, (C) R3, and (D) R4.

polymerization (R3 in Table 3, without GOx). As a reference, the fluorescence spectra in Nile Blue were initially recorded in various solvents (1.76 × 10−5 mol/L; Figure 6B), i.e., water and toluene (the solvents of the droplet and continuous phases, respectively) and toluene/t-BuOH/water (74.6/25/ 0.4, vol %) as this is the solvent mixture employed to study the enzyme activity (Figure 7). The fluorescence spectra of Nile Blue in the water droplets (i.e., as a miniemulsion) could not be recorded as the miniemulsion was too milky and strong scattering was observed even at high dilution. The bluish appearance of this solution however suggests that the dye is predominantly located in the dispersed aqueous phase (given that Nile Blue appears blue in water; Figure 6A,C). With the addition of t-BuOH, the solution immediately turned pink, which is indicative of a more hydrophobic environment for

Nile Blue (Figure 6A). The clear appearance also suggests similar refractive indices of the nanocapsule core, the shell, and the continuous phase. The emission maximum of Nile Blue was in agreement with the maximum found in the toluene/tBuOH/water mixture (Figure 6B). Fluorescence spectroscopic analysis of the continuous phase after removing the NB-loaded F

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Figure 7 shows the relative GOx activity in PLMA-based (R1 and R2) and PtBMA-based (R3 and R4) nanocapsules. All crude samples, which contained some unreacted monomer, showed good activity although R4 displayed a somewhat lower activity than the others. Because our assay is measuring relative activity, the high activity of the capsules does not necessarily mean no denaturation of the enzyme occurs. Confinement of the enzyme, solvent composition, and diffusion rates of substrate and product across the capsule membrane will all affect the enzyme activity. However, the diffusion of t-BuOH into PBS core of the capsules is expected to have little effect on enzyme activity (Figure S5), and the fact that the overall activity of the capsules is high relative to the free enzyme control suggests good protection of the enzyme by this method and good permeability of the substrate through the membrane. The type of the polymeric shell, despite significant differences in Tg of the base polymer (see above), did not exert any significant effect on the GOx activity. R2 and R4 were synthesized using a higher monomer and cross-linker loading than R1 and R3, and therefore R2 and R4 had thicker shells than R1 and R3.29 Interestingly, the shell thickness (Table 3) did not have any significant effect on enzyme activity either. The nanocapsules can protect GOx from organic solvents while allowing sufficient diffusion of substrates into the nanocapsules regardless of shell type or shell thickness as explored in this work. When the nanocapsules were precipitated in methanol to remove unreacted monomer and initiator, and subsequently redispersed in toluene, the encapsulated enzyme lost all activity (Figure 7). This is most likely caused by methanol to some extent entering the aqueous core, leading to enzyme denaturation.39 As a result, alternative ways were explored to purify and isolate the nanocapsules. The PLMA based nanocapsules (R1 and R2) were purified by centrifugation (14 500 rpm, 30 min) since PLMA is soluble in almost all nonpolar solvents. In contrast, the PtBMA based nanocapsules (R3 and R4) were purified by precipitation in nhexane, which is immiscible with water and thus unable to enter the core. All nanocapsules purified in these ways showed high activity (71−101% relative to free GOx in PBS/t-BuOH), although centrifugation or n-hexane precipitation led to somewhat lower activity than in crude nanocapsules. However, it should be noted that even the lowest activity measured was extremely high relative to unprotected enzyme in toluene, which showed almost no activity (Figure S5). The above results demonstrate that these nanocapsules can enable the use of enzymes such as GOx in entirely nonpolar organic solvents such as toluene. Enzyme-loaded nanocapsules have often been studied in water mixtures of polar solvents such as acetonitrile, THF, and dioxane, but very few methods are available for enabling their use in entirely organic media.11−13 Moreover, the encapsulation of enzymes in these polymer nanocapsules allows for easy separation of enzymes from reactants.

Figure 7. GOx activity in polymeric nanocapsules. “Crude”: crude nanocapsules with unreacted monomer remaining; “MeOH”: nanocapsules purified by precipitation in MeOH and centrifugation; “Cent/Hex”: nanocapsules purified by centrifugation only (R1 and R2) and precipitation with hexane followed by centrifugation (R3 and R4). GOx activity is a relative activity to native GOx in PBS (no encapsulation).

nanocapsules by centrifugation (14 500 rpm, 30 min) revealed the presence of Nile Blue, demonstrating that the polymeric nanocapsules synthesized using recipe R3 (Table 3) exhibit high permeability for small molecules, allowing rapid diffusion into the continuous phase in the present case. As such, it can be speculated that a small molecule substrate such as glucose can diffuse in and out through the shell. Measurement of GOx Activity. GOx is a widely used enzyme in biotechnology as well as the food industry, employed to determine the concentration of glucose in body fluids or food products. In addition, GOx has various applications such as a degassing agent and antibacterial agent.38 GOx, like many other enzymes and proteins, is prone to denaturation when exposed to heat and organic solvents such as toluene when not protected (Figure S5). We observed that heating the enzyme at 60 °C for 2 h resulted in complete deactivation of GOx (Figure S3A). The effect of ultrasonication and light irradiation on GOx activity was also separately investigated (Figure S3). The ultrasonication and visible light irradiation conditions used to encapsulate the enzyme via the IMEPP process were sufficiently mild to avoid denaturation (damaging shearing in the case of ultrasonication) of the enzyme, as revealed by the negligible effect on GOx activity in PBS (Figure S3). The activity of encapsulated GOx was determined by measuring the amount of hydrogen peroxide formed in a given time with a modified HRP/TMB assay. Because native GOx is inactive in the toluene/water mixture (Figure S5), the activity of the encapsulated GOx was measured relative to a standard curve of GOx in PBS. As glucose is insoluble in toluene, it was added as a solution in t-BuOH, which enabled diffusion to the interior of the nanocapsules. After 10 min of incubation with the glucose, the concentration of generated H2O2 was measured by first diluting the particles in t-BuOH (to extract all of the formed H2O2 from the inside of the particles) and then adding this solution to a solution of HRP and TMB. A large excess of these reagents was used so that all of the available H2O2 was consumed in less than 1 min (Figure S6). The amount of H2O2 generated by the encapsulated GOx was compared to the standard curve of unmodified GOx in PBS (Figure S4).

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CONCLUSIONS GOx-loaded polymer nanocapsules have been successfully synthesized via the IMEPP technique under blue light irradiation employing photoinitiation. The use of light irradiation for the polymerization instead of thermal initiation was found to be the key to maintaining high activity of the encapsulated enzyme. Having identified the optimal purification conditions, the encapsulated GOx retained almost all of its activity in an organic solvent environment that would G

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biocatalyst for use in organic media. Biotechnol. Technol. 1989, 3 (4), 275−280. (12) Daubresse, C.; Grandfils, C.; Jerome, R.; Teyssie, P. Enzyme Immobilization in Nanoparticles Produced by Inverse Microemulsion Polymerization. J. Colloid Interface Sci. 1994, 168 (1), 222−229. (13) Kobayashi, J.; Mori, Y.; Kobayashi, S. Novel immobilization method of enzymes using a hydrophilic polymer support. Chem. Commun. 2006, 40, 4227−4229. (14) Vrignaud, S.; Benoit, J.-P.; Saulnier, P. Strategies for the nanoencapsulation of hydrophilic molecules in polymer-based nanoparticles. Biomaterials 2011, 32 (33), 8593−8604. (15) Ulu, A.; Ates, B. Immobilization of l-Asparaginase on Carrier Materials: A Comprehensive Review. Bioconjugate Chem. 2017, 28 (6), 1598−1610. (16) Jiskoot, W.; Randolph, T. W.; Volkin, D. B.; Middaugh, C. R.; Schöneich, C.; Winter, G.; Friess, W.; Crommelin, D. J. A.; Carpenter, J. F. Protein instability and immunogenicity: Roadblocks to clinical application of injectable protein delivery systems for sustained release. J. Pharm. Sci. 2012, 101 (3), 946−954. (17) Yan, M.; Ge, J.; Liu, Z.; Ouyang, P. Encapsulation of Single Enzyme in Nanogel with Enhanced Biocatalytic Activity and Stability. J. Am. Chem. Soc. 2006, 128 (34), 11008−11009. (18) Wei, W.; Du, J.; Li, J.; Yan, M.; Zhu, Q.; Jin, X.; Zhu, X.; Hu, Z.; Tang, Y.; Lu, Y. Construction of Robust Enzyme Nanocapsules for Effective Organophosphate Decontamination, Detoxification, and Protection. Adv. Mater. 2013, 25 (15), 2212−2218. (19) Liu, Y.; Du, J.; Yan, M.; Lau, M. Y.; Hu, J.; Han, H.; Yang, O. O.; Liang, S.; Wei, W.; Wang, H.; Li, J.; Zhu, X.; Shi, L.; Chen, W.; Ji, C.; Lu, Y. Biomimetic enzyme nanocomplexes and their use as antidotes and preventive measures for alcohol intoxication. Nat. Nanotechnol. 2013, 8 (3), 187−192. (20) Du, J.; Jin, J.; Liu, Y.; Li, J.; Tokatlian, T.; Lu, Z.; Segura, T.; Yuan, X.-b.; Yang, X.; Lu, Y. Gold-Nanocrystal-Enhanced Bioluminescent Nanocapsules. ACS Nano 2014, 8 (10), 9964−9969. (21) Gu, Z.; Dang, T. T.; Ma, M.; Tang, B. C.; Cheng, H.; Jiang, S.; Dong, Y.; Zhang, Y.; Anderson, D. G. Glucose-Responsive Microgels Integrated with Enzyme Nanocapsules for Closed-Loop Insulin Delivery. ACS Nano 2013, 7 (8), 6758−6766. (22) Kim, J.; Grate, J. W.; Wang, P. Nanostructures for enzyme stabilization. Chem. Eng. Sci. 2006, 61 (3), 1017−1026. (23) Kim, J. B.; Grate, J. W.; Wang, P. Nanobiocatalysis and its potential applications. Trends Biotechnol. 2008, 26 (11), 639−646. (24) Silva, C.; Martins, M.; Jing, S.; Fu, J.; Cavaco-Paulo, A. Practical insights on enzyme stabilization. Crit. Rev. Biotechnol. 2017, 1−16. (25) Utama, R. H.; Drechsler, M.; Förster, S.; Zetterlund, P. B.; Stenzel, M. H. Synthesis of pH-Responsive Nanocapsules via Inverse Miniemulsion Periphery RAFT Polymerization and Post-Polymerization Reaction. ACS Macro Lett. 2014, 3, 935−939. (26) Utama, R. H.; Jiang, Y.; Zetterlund, P. B.; Stenzel, M. H. Biocompatible Glycopolymer Nanocapsules via Inverse Miniemulsion Periphery RAFT Polymerization for the Delivery of Gemcitabine. Biomacromolecules 2015, 16 (7), 2144−2156. (27) Utama, R. H.; Guo, Y.; Zetterlund, P. B.; Stenzel, M. H. Synthesis of hollow polymeric nanoparticles for protein delivery via inverse miniemulsion periphery RAFT polymerization. Chem. Commun. 2012, 48, 11103−5. (28) Ishizuka, F.; Utama, R. H.; Kim, S.; Stenzel, M. H.; Zetterlund, P. B. RAFT inverse miniemulsion periphery polymerization in binary solvent mixtures for synthesis of nanocapsules. Eur. Polym. J. 2015, 73, 324−334. (29) Utama, R. H.; Dulle, M.; Forster, S.; Stenzel, M. H.; Zetterlund, P. B. SAXS Analysis of Shell Formation During Nanocapsule Synthesis via Inverse Miniemulsion Periphery RAFT Polymerization. Macromol. Rapid Commun. 2015, 36 (13), 1267−71. (30) Utama, R. H.; Stenzel, M. H.; Zetterlund, P. B. Inverse Miniemulsion Periphery RAFT Polymerization: A Convenient Route to Hollow Polymeric Nanoparticles with an Aqueous Core. Macromolecules 2013, 46, 2118−2127.

otherwise lead to total loss of activity through denaturation. The methodology developed is versatile as the guest molecule can be easily replaced with other enzymes as well as any other water-soluble compound. It can be applied to the encapsulation of various enzymes as biocatalysts and enable them to function in nonpolar organic solvents. In addition, this approach has great potential for generation of nanocarriers for therapeutic protein applications.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.macromol.7b02377.

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Experimental details, theoretical calculation, additional TEM images (PDF)

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected] (M.H.S.). *E-mail: [email protected] (P.B.Z.). ORCID

Per B. Zetterlund: 0000-0003-3149-4464 Martina H. Stenzel: 0000-0002-6433-4419 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors thank the Australian Research Council (ARC DP170101191) and the University of New South Wales for funding.

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REFERENCES

(1) Iyer, P. V.; Ananthanarayan, L. Enzyme stability and stabilizationAqueous and non-aqueous environment. Process Biochem. 2008, 43 (10), 1019−1032. (2) Sheldon, R. A.; van Pelt, S. Enzyme immobilisation in biocatalysis: why, what and how. Chem. Soc. Rev. 2013, 42 (15), 6223−6235. (3) Hasan, F.; Shah, A. A.; Hameed, A. Industrial applications of microbial lipases. Enzyme Microb. Technol. 2006, 39 (2), 235−251. (4) Bornscheuer, U. T.; Huisman, G. W.; Kazlauskas, R. J.; Lutz, S.; Moore, J. C.; Robins, K. Engineering the third wave of biocatalysis. Nature 2012, 485 (7397), 185−194. (5) Klibanov, A. M. Why are enzymes less active in organic solvents than in water? Trends Biotechnol. 1997, 15 (3), 97−101. (6) Carrea, G.; Riva, S. Properties and Synthetic Applications of Enzymes in Organic Solvents. Angew. Chem., Int. Ed. 2000, 39 (13), 2226−2254. (7) Stepankova, V.; Bidmanova, S.; Koudelakova, T.; Prokop, Z.; Chaloupkova, R.; Damborsky, J. Strategies for Stabilization of Enzymes in Organic Solvents. ACS Catal. 2013, 3 (12), 2823−2836. (8) Veronese, F. M. Peptide and protein PEGylation: a review of problems and solutions. Biomaterials 2001, 22 (5), 405−417. (9) Roberts, M. J.; Bentley, M. D.; Harris, J. M. Chemistry for peptide and protein PEGylation. Adv. Drug Delivery Rev. 2012, 64, 116−127. (10) Xu, D. Y.; Yang, Y.; Yang, Z. Activity and stability of crosslinked tyrosinase aggregates in aqueous and nonaqueous media. J. Biotechnol. 2011, 152 (1), 30−36. (11) Khmelnitsky, Y. L.; Neverova, I. N.; Momtcheva, R.; Yaropolov, A. I.; Belova, A. B.; Levashov, A. V.; Martinek, K. Surface-modified polymeric nanogranules containing entrapped enzymes: A novel H

DOI: 10.1021/acs.macromol.7b02377 Macromolecules XXXX, XXX, XXX−XXX

Article

Macromolecules (31) Stenzel, M. H.; Barner-Kowollik, C. The living dead - common misconceptions about reversible deactivation radical polymerization. Mater. Horiz. 2016, 3 (6), 471−477. (32) Mclaren, A. D.; Luse, R. A. Mechanism of Inactivation of Enzyme Proteins by Ultraviolet Light. Science 1961, 134 (3482), 836−837. (33) Lante, A.; Tinello, F.; Lomolino, G. Effect of UV light on microbial proteases: From enzyme inactivation to antioxidant mitigation. Innovative Food Sci. Emerging Technol. 2013, 17, 130−134. (34) Mitsukami, Y.; Donovan, M. S.; Lowe, A. B.; McCormick, C. L. Water-Soluble Polymers. 81. Direct Synthesis of Hydrophilic StyrenicBased Homopolymers and Block Copolymers in Aqueous Solution via RAFT. Macromolecules 2001, 34, 2248−2256. (35) Pasquali, R. C.; Taurozzi, M. P.; Bregni, C. Some considerations about the hydrophilic−lipophilic balance system. Int. J. Pharm. 2008, 356 (1), 44−51. (36) Brandrup, J.; Immergut, E. H.; Grulke, E. A. Polymer Handbook, 4th ed.; John Wiley & Sons: New York, 1975. (37) Libertino, S.; Aiello, V.; Scandurra, A.; Renis, M.; Sinatra, F. Immobilization of the Enzyme Glucose Oxidase on Both Bulk and Porous SiO2 Surfaces. Sensors 2008, 8 (9), 5637−5648. (38) Wong, C. M.; Wong, K. H.; Chen, X. D. Glucose oxidase: natural occurrence, function, properties and industrial applications. Appl. Microbiol. Biotechnol. 2008, 78 (6), 927−938. (39) Chapman, R.; Gormley, A. J.; Herpoldt, K.-L.; Stevens, M. M. Highly Controlled Open Vessel RAFT Polymerizations by Enzyme Degassing. Macromolecules 2014, 47 (24), 8541−8547.

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DOI: 10.1021/acs.macromol.7b02377 Macromolecules XXXX, XXX, XXX−XXX