Stabilization of Lipase in Polymerized High Internal Phase Emulsions

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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.

ACS Paragon Plus Environment

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

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1306.

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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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a Cross-Linkable Core. Methods Enzymol. 2016, 571, 1–17.

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(39) Talbert, J. N.; Wang, L.-S.; Duncan, B.; Jeong, Y.; Andler, S. M.; Rotello, V. M.; Goddard,

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J. M. Immobilization and Stabilization of Lipase (CaLB) through Hierarchical Interfacial

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


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(40) Andler, S. M.; Wang, L.-S.; Rotello, V. M.; Goddard, J. M. Influence of Hierarchical

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(41) Mol, J. C. Industrial Applications of Olefin Metathesis. J. Mol. Catal. Chem. 2004, 213 (1), 39–45.

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(42) Handbook of Metathesis; Grubbs, R. H., Ed.; Wiley-VCH: Weinheim, 2003.

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(43) Waghmare, G. V.; Rathod, V. K. Ultrasound Assisted Enzyme Catalyzed Hydrolysis of

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

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FIGURE CAPTIONS

336

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

337

the polymerization of the monomer.

338

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

339

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.

342

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

343

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

344

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

345

deviation.

346

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

347

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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349

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

350

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

351

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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Stabilization of Lipase in Polymerized High Internal Phase Emulsions

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