Stabilization of Lipase in Polymerized High Internal Phase Emulsions

Mar 27, 2018 - Candida antarctica lipase B is stabilized in a porous, high internal phase emulsion (HIPE) of polydicyclopentadiene to enable biocataly...
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Stabilization of lipase in polymerized high internal phase emulsions Stephanie M Andler, and Julie M Goddard J. Agric. Food Chem., Just Accepted Manuscript • Publication Date (Web): 27 Mar 2018 Downloaded from http://pubs.acs.org on March 27, 2018

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Journal of Agricultural and Food Chemistry

Stabilization of lipase in polymerized high internal phase emulsions Stephanie M. Andler and Julie M. Goddard* Department of Food Science, Cornell University, Ithaca, NY 14853.

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ABSTRACT

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Candida antarctica lipase B is stabilized in a porous, high internal phase emulsion (HIPE) of

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polydicyclopentadiene to enable biocatalytic waste stream upcycling. The immobilized lipase is

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subjected to thorough washing conditions and tested for stability in extreme environments and

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reusability. A porous internal microstructure is revealed through scanning electron microscopy.

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After preparation, lipase activity increased to 139±9.7% of its original activity. After ten cycles

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of reuse, immobilized lipase retains over 50% activity. Immobilized lipase retains activity after

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24 hours exposure to temperatures ranging from 20 ˚C to 60 ˚C, and pH values of 3, 7, and 10. In

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the most extreme environments tested, lipase retained 42.8±21 % relative activity after exposure

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to 60 ˚C, and 49.4±16% relative activity after exposure to pH 3. Polymerized HIPEs stabilize

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lipase, and thus extend its working range. Further synthesis optimization has the potential to

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increase enzyme stability, immobilization efficiency, and uniformity. The reported hierarchical

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stabilization technique shows promise for use of immobilized lipase in non-ideal, industrially

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relevant conditions.

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KEYWORDS: lipase, immobilization, enzyme, HIPE

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Journal of Agricultural and Food Chemistry

INTRODUCTION

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Industrial food processing waste streams represent a significant environmental burden, as an

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estimated 30-40% of all food is wasted.1–3 Enzymes have the potential to increase sustainability

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through the valorization of waste streams. Specifically, lipase B from Candida antarctica has

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been extensively studied for its ability to catalyze a variety of reactions with high specificity. 4,5

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For example, lipase can transform waste oil streams through esterification reactions,6 an important

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example of waste stream upcycling, as over 2 billion pounds of yellow grease were produced in

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the United States in 2015 alone.7 Lipase has been explored as a biocatalyst for biodiesel production

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from waste oil because of its specificity, lack of soap formation, and performance in the presence

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of interfering substances (e.g. free fatty acids, water).8–12 However, these esterification reactions

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typically require non-ideal conditions for native lipase (e.g. increased temperatures), which

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decrease activity and consequently reduce overall product conversion.13,14 Additionally, the cost

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of enzymes can be prohibitive to commercial adoption.

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To address these hurdles to commercialization, researchers have explored enzyme

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immobilization.15–17 The main methods to immobilize enzymes include binding to a carrier,

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entrapment and encapsulation, and cross-linking.17 The properties of the support material can

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influence the stability and activity of the enzyme; for example, pore size can impact diffusion of

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the substrate, thus influencing activity.18 Further, while in many cases a hydrophilic support is

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favored in retaining immobilized enzyme activity, many lipases are reported to undergo interfacial

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activation, in which the enzyme assumes an open, more active, form when situated at a

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hydrophobic interface.19–22 While not all sources of lipase exhibit interfacial activation,23–25

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hydrophobic materials are of interest for lipase immobilization.

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When considering the size and morphology of the solid support to which enzymes are

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immobilized, enzyme immobilization research has focused on either macroscale or nanoscale

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materials.15,26 While immobilization onto macroscale supports enables facile recovery, nanoscale

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immobilization can allow the enzyme to experience conditions similar to the native enzyme.

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However, as the size of the support decreases, the time for particle recovery increases.27 Thus,

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hierarchical structures, leveraging both nano- and macroscale materials, have gained interest

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because of their ability to leverage attributes from both orders of scale.26,28–30 Highly porous

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materials of appropriate pore size can present increased surface area for immobilization.18

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Importantly, these porous materials can be formed at macroscale sizes to permit facile handling

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and recovery.

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High internal phase emulsions (HIPEs) are porous structures, defined as having greater than 70%

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internal phase volume.31 Polymerized HIPEs have been produced from a variety of monomers

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including styrene, acrylates, methacrylates, and dicyclopentadiene.32–35 Few examples exist of

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enzyme containing HIPEs, which immobilize the enzyme after HIPE formation. 36,37 To our

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knowledge, no research exists on the incorporation of enzymes into the HIPE during

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polymerization.

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We have previously prepared cross-linked microparticles of polydicyclopentadiene with

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interfacially assembled nanoparticle-conjugated lipase.38–40

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mechanically robust polymer, produced from a ring opening metathesis polymerization.41,42 These

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~10 µM particles showed enhanced lipase stability across a wide range of pH values and

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temperatures, but were difficult to recover from viscous solvents.40 As a result, the present work

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synthesized lipase-containing macroscale polydicyclopentadiene HIPEs.

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Materials and Methods

Polydicyclopentadiene is a

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Materials

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Candida antarctica lipase B was kindly donated from c-LEcta (Leipzig, Germany). Resorufin

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butyrate was purchased from Santa Cruz Biotechnology (Dallas, TX, USA). Dicyclopentadiene,

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dimethylformamide (DMF), and toluene were purchased from Fisher Scientific (Fairlawn, NJ,

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USA). First generation Grubbs’ catalyst and Pluronic® L121 (poly(ethylene glycol)-block-

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poly(propylene glycol)-block-poly(ethylene glycol)) were purchased from Sigma-Aldrich (Saint

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Louis, MO, USA).

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High Internal Phase Emulsion (HIPE) Preparation

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HIPEs were fabricated using previously described methods, with modifications.34,35 The HIPE

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was prepared as 80% aqueous phase and 20% oil phase (v/v). Of the oil phase, 90% was Pluronic®

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L121 in dicyclopentadiene monomer (1% vol Pluronic® L121/vol monomer), and 10% Grubbs’

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catalyst (first generation) in toluene (5% wt Grubbs’ catalyst/vol toluene). For the aqueous phase,

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lipase was dissolved in 10 mM 2-(N-morpholino)ethanesulfonic acid (MES) buffer, pH 7 at a

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concentration of 50 µg protein/ml. Grubbs’ catalyst (first generation) was dissolved in toluene by

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sonicating for 10 minutes. Pluronic® L121 and dicyclopentadiene were blended for 5 minutes

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using magnetic stirring. Then, for the lipase containing samples, the lipase in MES buffer solution

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was added dropwise over 5 minutes. For the control HIPEs, 10 mM MES pH 7 was added

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dropwise over 5 minutes. The emulsion was then stirred for an additional two minutes. The

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solution of Grubbs’ catalyst was then added, and the emulsion was stirred for an additional two

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minutes. The emulsion was then poured into a three-cavity Teflon mold (21x 120mm, Ted Pella,

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Redding, CA, USA), and covered with embedding film (ACLAR Embedding Film 33C, Ted Pella,

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Redding, CA, USA). Embedding film covers were utilized to prevent shrinkage, which can occur

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upon rapid evaporation of the monomer.34

Polymerization was performed under ambient

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conditions (20 ˚C) to prevent denaturation of lipase. After 5 hours polymerization at 20 ˚C, the

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embedding film was removed, and HIPEs were dried overnight in the fume hood. The HIPEs

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were cut into 10 pieces per cavity, weighing approximately 30 mg each. HIPEs were washed in

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10 mM MES buffer, pH 7 for 5 cycles of 30 minutes at 20 ˚C.

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HIPE Activity

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HIPE activity was determined using the synthetic substrate 10 µM resorufin butyrate in pre-

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warmed 10 mM MES buffer pH 7, 50 ˚C. Resorufin butyrate was first dissolved in DMF at a

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concentration of 5.19 mM, then diluted to a final concentration of 10 µM in prewarmed 10 mM

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MES buffer pH 7. A piece of HIPE (approximately 30 mg) was placed in a 2 ml microcentrifuge

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tube. 600 µl of prewarmed substrate was added to the tube, and quickly vortexed before placing

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in a 50 ˚C incubator. The HIPEs were incubated for 3 minutes, then 300 µl was transferred to a

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black, 96-well plate. The samples were read using a plate reader (Synergy Neo 2, Biotek,

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Winooski, VT, USA) at 528/590 nm excitation and emission wavelengths, respectively. As a

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control, HIPEs prepared without lipase were tested for activity using the same procedure, and the

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fluorescence was subtracted from the HIPEs containing lipase.

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HIPE Washing Activity

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Freshly prepared HIPEs were washed for 5 cycles of 30 minutes at 20 ˚C in 10 mM MES buffer,

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pH 7. HIPEs were washed in a 50 ml centrifuge tube at a ratio of 1 mL buffer per ~30mg piece of

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HIPE. Tubes were rotated (Rotating Mixer R5010, Benchmark Scientific, Sayreville, NJ, USA)

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at 30 RPM. Following each wash cycle, four HIPEs were removed from the washing buffer, and

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tested for activity using the procedure described previously.

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HIPE Reuse

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Reuse of the HIPEs was determined by repeated exposure to the substrate, resorufin butyrate, and

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subsequent activity determination. Control and lipase containing HIPEs were tested for activity

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using the procedure described previously. After each exposure to the substrate, the HIPEs were

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washed 2 x 30 sec using 10 mM MES buffer, pH 7, then exposed to the substrate again. The

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process was repeated for a total of 10 cycles. Activity of subsequent cycles was compared relative

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to the activity of the first cycle.

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HIPE Thermostability

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Control and lipase containing HIPEs were first tested for activity using the procedure outlined

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previously. The HIPEs were then washed 2 x 30 sec in 10 mM MES buffer, pH7, then incubated

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at 20 ˚C, 40 ˚C, 50 ˚C, and 60 ˚C in 600 µl 10 mM MES buffer pH 7 for 24 hours. After 24 hours,

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the storage buffer was removed, and HIPE activity was tested according to the procedure outlined

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previously.

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incubation.

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HIPE pH Stability

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Control and lipase containing HIPEs were first tested for activity using the procedure outlined

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previously. The HIPEs were then washed 2 x 30 sec in 10 mM MES buffer, pH7, and incubated

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at pH 3, 7, and 10 in 600 µl 10 mM citrate, phosphate, and carbonate buffers, respectively, for 24

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hours at 20 ˚C. After 24 hours, the storage buffer was removed and HIPEs were washed with 0.1

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M MES buffer, pH 7. Then, activity was tested according to the procedure outlined previously.

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Activity of each HIPE was compared to its starting activity prior to incubation.

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Scanning Electron Microscopy

Activity of each HIPE was compared to its individual starting activity prior to

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Scanning electron micrographs were obtained using a benchtop SEM (JEOL Neoscope JCM 6000)

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operating at 15 kV (Nikon Instruments, Inc. Melville, NY).

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(Cressington Sputter Coater 108auto, Watford, UK) with gold prior to analysis. The reported

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image is representative of 9 images acquired across each of two independently prepared samples.

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Statistical Analysis

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All experiments were performed using n=4 samples, and repeated on a second day, using

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independently prepared HIPEs. HIPEs were synthesized in the absence of lipase and used as a

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negative control for all experiments. Statistical analysis was performed using Graphpad

Samples were sputter coated

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Prism software (v. 7.01, GraphPad Software, La Jolla, CA). All experimental data were treated

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as paired data. For Washing Activity, Thermostability, and pH Stability experiments, one-way

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analysis of variance (ANOVA) with Tukey’s pairwise comparison for the identification of

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statistical differences was performed using p < 0.05. For Reuse experiments, One-way analysis

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of variance (ANOVA) with Dunnet’s pairwise comparison for the identification of statistical

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differences was performed using p < 0.05.

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RESULTS AND DISCUSSION

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HIPE synthesis overcomes some hurdles to commercial adoption of immobilized enzymes, as

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only mixers and casting molds are needed, and synthesis can be readily scaled up. Scanning

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electron microscopy (SEM) (Figure 1) revealed a porous internal microstructure, confirming

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successful formation of the polymerized HIPEs and the presence of a hierarchical structure. This

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porosity provided a high surface area amenable for protein loading. Initial activity of as-prepared

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HIPEs revealed approximately 139±9.7% activity retention compared to native lipase. However,

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lipase was not strongly immobilized, as shown by a decrease in relative activity (as compared to

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as-prepared HIPE activity) with each subsequent wash cycle (Figure 2). Following an initial

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decrease, relative activity stabilized after the fourth wash, representing activity of immobilized

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lipase. The observed decrease in activity suggests loss of lipase from weak immobilization,

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possibly a result of the surface-active nature of the surfactant, which may have competed with

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lipase for sites in interfacial assembly. To ensure subsequent tests evaluated only immobilized

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lipase, stability and reuse experiments used HIPEs that had undergone the 5-cycle washing step.

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Reported values represent the mean of n=4 samples, with error bars representing standard

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deviation, and are representative of experiments repeated on two days, with independently

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prepared HIPEs.

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The performance of the immobilized lipase HIPEs under conditions of pH and temperature

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extremes was determined to characterize efficacy under industrially relevant conditions. To

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determine HIPE stability in varying pH environments, HIPEs were tested for initial activity, then

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washed and stored in pH 3, 7, and 10 buffers for 24 hours at 20 ˚C (Figure 3). After storage at pH

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3, HIPEs retained 49.4±16% activity. This activity retention is an improvement over native lipase,

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which has been shown to lose almost all activity at pH 3.39 At pH 7, HIPEs retained 76.7±7%

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activity, suggesting favorable working conditions at neutral pH. At pH 10, HIPEs retained

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62.7±5% activity, showing that immobilization within polydicyclopentadiene broadens the

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working range of lipase. We hypothesize that the polydicyclopentadiene is providing stability to

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the structure of lipase, and possibly influences the microenvironment that lipase experiences. 40

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The observed decrease in activity across all pH values may also be attributed to weak

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immobilization and migration of the enzyme.

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To determine thermostability of immobilized lipase, HIPEs were tested for activity, washed, and

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stored in 10 mM MES buffer (pH 7) at 20 ˚C, 40 ˚C, 50 ˚C, and 60 ˚C for 24 hours. After exposure,

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HIPEs retained 111.6±23, 157.5±21, 54.0±19, 42.8±21 % activity at 20 ˚C, 40 ˚C, 50 ˚C, and 60

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˚C, respectively (Figure 4). At 20 ˚C and 40 ˚C, HIPEs showed an increase in activity compared

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to their starting activity. These data further support that stabilizing interactions occur between

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polydicyclopentadiene and lipase. HIPEs retained activity after storage at 50 ˚C and 60 ˚C, in

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contrast to native lipase which has been reported to lose all activity at elevated temperatures within

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approximately 2 hours, and Novozym 435, which loses greater than 60% activity at 60 ˚C.39

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Elevated temperatures are commonly required for esterification reactions, supporting the need for

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stabilized enzymes. These data further support the use of polydicyclopentadiene as a support

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material for lipase in extreme environments.

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Because enzymes are more expensive than their chemical counterparts, reusability is imperative

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for commercial adoption. For proof of concept, HIPEs were exposed to the resorufin butyrate for

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total of 10 cycles, with washes (10 mM MES buffer, pH 7) between cycles. Activity was

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monitored after each cycle and compared to the starting activity.

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activity after each reuse cycle. HIPEs retained over 50% activity after 10 cycles, showing promise

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for reusability.

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significant, could point towards improvement in the post-polymerization washing process.

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Additionally, concentrations of both protein and surfactant can be optimized to support lipase

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interfacial assembly. These results are in agreement with practical application of Novozym 435

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to hydrolysis of waste cooking oil, where 62% activity remained after 4 cycles of reuse. 43

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Nevertheless, the sustained activity over 10 cycles suggests potential for reuse.

Figure 5 shows average HIPE

The initial observed drop in activity (63.9±9.5%), while not statistically

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These results show the potential for use of hierarchical materials as solid supports in the

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stabilization of enzymes for industrial waste reutilization processes. Immobilized lipase was

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stabilized in polymerized HIPEs and retained activity after exposure to non-ideal environments,

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retaining approximately 49% activity at pH 3 and approximately 42% activity at 60 ˚C after 24

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hours exposure. Further research is necessary to determine stability in practical solvents and

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further in practical applications; however, the tailorable size of the HIPE makes the process

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suitable for scale-up and customization. HIPE synthesis optimization may further improve lipase

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stability; for example, manipulating protein concentration, mixing shear, aqueous phase percent,

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and surfactant type.

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immobilization and offer advantages over free enzymes. Enzymatic valorization of waste streams

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can eliminate the need for toxic solvents and reduce the formation of by-products.

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optimization, lipases immobilized in polymerized HIPEs can be considered for industrial

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esterification reactions. Further, the value-added reutilization of waste streams, and greener

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processing methods can help improve the environmental sustainability of industrial food

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manufacturing.

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AUTHOR INFORMATION

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Corresponding Author

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*[email protected]. * Department of Food Science, Cornell University, Ithaca, NY 14853

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Author Contributions

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The manuscript was written through contributions of all authors. All authors have given approval

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to the final version of the manuscript.

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Notes

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There are no conflicts to declare.

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ACKNOWLEDGMENT

Nevertheless, hierarchical structures show promise for enzyme

With further

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This project was supported, in part, by the USDA Agriculture and Food Research Initiative Grant

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no. 2014-67021-21584, and the Cornell Institute for Food Systems Industry Partnership Program.

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ABBREVIATIONS

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HIPE, high internal phase emulsion; MES, 2-(N-morpholino)ethanesulfonic acid; SEM, scanning electron microscopy.

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Approach to Enzyme Stabilization and Immobilization. Enzyme Microb. Technol. 2006, 39

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(29) Cipolatti, E. P.; Silva, M. J. A.; Klein, M.; Feddern, V.; Feltes, M. M. C.; Oliveira, J. V.;

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Ninow, J. L.; de Oliveira, D. Current Status and Trends in Enzymatic Nanoimmobilization.

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J. Mol. Catal. B Enzym. 2014, 99, 56–67.

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(31) Lissant, K. Emulsions and Emulsion Technology; CRC Press, 1974; Vol. 3.

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(32) Silverstein, M. S. PolyHIPEs: Recent Advances in Emulsion-Templated Porous Polymers.

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Prog. Polym. Sci. 2014, 39 (1), 199–234.

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(33) Kovačič, S.; Matsko, N. B.; Jerabek, K.; Krajnc, P.; Slugovc, C. On the Mechanical

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Properties of HIPE Templated Macroporous Poly(Dicyclopentadiene) Prepared with Low

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Surfactant Amounts. J Mater Chem A 2013, 1 (3), 487–490.

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(34) Kovačič, S.; Preishuber-Pflügl, F.; Slugovc, C. Macroporous Polyolefin Membranes from

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Dicyclopentadiene High Internal Phase Emulsions: Preparation and Morphology Tuning.

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Macromol. Mater. Eng. 2014, 299 (7), 843–850.

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(35) Kovačič, S.; Matsko, N. B.; Ferk, G.; Slugovc, C. Macroporous Poly(Dicyclopentadiene)

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ΓFe2O3/Fe3O4 Nanocomposite Foams by High Internal Phase Emulsion Templating. J.

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Mater. Chem. A 2013, 1 (27), 7971.

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Carbonaceous Foams Electrodes for Biofuel Cells. Energy Environ. Sci. 2010, 3 (9), 1302–

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Lipase Immobilized onto Hydrophobic Microporous Styrene–divinylbenzene Copolymer.

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Microparticles by Interfacial Self-Assembly of Enzyme–Nanoparticle Conjugates Around

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Assembly. Biomacromolecules 2014, 15 (11), 3915–3922.

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Interfacial Assembly on Lipase Stability and Performance in Deep Eutectic Solvent. J.

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Waste Cooking Oil under Solvent Free Condition. Ultrason. Sonochem. 2016, 32, 60–67.

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FIGURE CAPTIONS

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Figure 1. SEM micrograph of cross-section of HIPE. A highly porous network is formed though

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the polymerization of the monomer.

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Figure 2. Relative percent activity of HIPEs after each 30-minute was with 10 mM MES buffer,

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pH 7. Relative activity was determined using resorufin butyrate and was calculated by comparison

340

to the starting activity of unwashed HIPEs. Values represent means of n=4 determinations with

341

error bars representing standard deviation.

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Figure 3. Activity of HIPEs after exposure to pH 3, 7, 10 for 24 hours. Relative activity was

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determined using resorufin butyrate and compared to the starting activity of each HIPE after

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washing. Values represent means of n=4 determinations with error bars representing standard

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deviation.

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Figure 4. Activity of HIPEs after 24 hours exposure to 20 ˚C, 40 ˚C, 50 ˚C, and 60 ˚C. Activity

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was determined using resorufin butyrate and compared to the starting activity of each HIPE.

348

Values represent means of n=4 determinations with error bars representing standard deviation.

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Figure 5. Activity of HIPEs after exposure to resorufin butyrate for a total of 10 cycles. Over

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50% activity is retained after 10 cycles of reuse. Values represent means of n=4 determinations

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with error bars representing standard deviation.

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FIGURE GRAPHICS Figure 1

Figure 2

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Figure 3

Figure 4

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Figure 5

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GRAPHIC FOR TABLE OF CONTENTS

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