- Email: [email protected]
Enzyme-initiated free radical polymerizations of vinyl monomers using horseradish peroxidase
CHAPTER NINE
Enzyme-initiated free radical polymerizations of vinyl monomers using horseradish peroxidase Kyle J. Rodrigueza, Michela M. Pellizzonia, Robert J. Chadwickb, Chao Guob, Nico Brunsa,b,* a
Adolphe Merkle Institute, University of Fribourg, Fribourg, Switzerland Department of Pure and Applied Chemistry, University of Strathclyde, Glasgow, United Kingdom *Corresponding author: e-mail address: [email protected] b
Contents 1. Introduction 2. Free radical polymerizations of vinyl monomers using horseradish peroxidase 2.1 Polymerizations 3. Polymer characterization 3.1 Nuclear magnetic resonance (NMR) spectroscopy 3.2 Gel permeation chromatography (GPC) 4. Conclusions Acknowledgments References
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Abstract In this chapter, we highlight the use of horseradish peroxidase (HRP) as a catalyst to initiate free radical polymerizations of vinyl monomers under benign reaction conditions. A variety of vinyl monomers, including 4-acryloylmorpholine (AM), 2-hydroxyethyl methacrylate (HEMA), and poly(ethylene glycol) methyl ether acrylate (PEGA) were polymerized. The enzyme converts exogenous hydrogen peroxide into a usable radical source, which when coupled with a β-diketone, yields a radical that initiates chain growth in the presence of monomers. The resulting polymers were characterized using nuclear magnetic resonance (NMR) spectroscopy and gel permeation chromatography (GPC). By using enzymatic free radical polymerizations, polymers can be generated in a sustainable, environmentally-friendly, and scalable fashion.
Methods in Enzymology, Volume 627 ISSN 0076-6879 https://doi.org/10.1016/bs.mie.2019.08.013
#
2019 Elsevier Inc. All rights reserved.
249
250
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1. Introduction Free radical polymerizations (FRPs) are used every year worldwide to manufacture almost 45% of synthetic polymers such as rubber and plastics using peroxy compounds, redox systems or azo initiators (Binder, 2019; Braun, 2009). From a historical perspective, FRP was first reported by Simon (1839) and later by Blyth and Hofmann (1845) in 1839 and 1845, respectively. They observed the formation of glass like solids starting from a styrene containing natural resin isolated from the wounded bark of Liquidambar orientalis in the presence of light. The concept of polymers and radicals were introduce in the same century (by Berzelius, Berthelot and Wurtz) even if the meaning and nature of these entities remained ambiguous until the reports of Gomberg (1897) and Staudinger (1920) at the beginning of last century. The mechanism of FRP was postulated by Flory (1936) when several synthetic polymers obtained by condensation of formaldehyde with phenol, urea and proteins (e.g., Bakelite, Pollopas and Galalit), as well as cellulose derivatives were already on the market (Mark, 1964). Nowadays, due to their simple reaction conditions, FRPs have been implemented for generating materials such as hydrogels, drug vehicles, and packaging. These applications have resulted in FRPs being implemented as a common synthetic tool for generating highly valuable materials. Additionally, in the last decades, new methodologies have been developed to control radical polymerizations (reversible-deactivation radical polymerization techniques such as nitroxide mediated polymerizations (NMP), atom transfer radical polymerizations (ATRP), and reversible-addition-fragmentation chain transfer (RAFT) polymerizations) but their application on industrial scale is still limited and FRPs remain the most widely used techniques due to the well establish and simple reaction conditions (Binder, 2019). Incorporating biocatalysis into traditional chemical processes allows for a more sustainable approach for generating materials necessary for the food and pharmaceutical industries where biocompatibility is necessary. Harnessing the power of biology can be a very powerful tool, e.g., to repurpose enzymes in perform non-native transformations, which result in efficient catalysis through biological mechanisms. After the first report in 1951 by Parravano, who reported the polymerization of methyl methacrylate in the presence of Xantine oxidase and formaldehyde (Parravano, 1951) many enzymes like oxidases and peroxidases were found to be able to catalyze radical polymerizations of a variety of monomers ranging from phenolic compounds to those containing vinyl
Enzyme-initiated free radical polymerizations
251
groups (Derango, Chiang, Dowbenko, & Lasch, 1992; Hollmann, 2010; Hollmann & Arends, 2012; Singh, Ma, & Kaplan, 2000; Zavada, Battsengel, & Scott, 2016). The use of biocatalysts to initiate and/or mediate polymerizations not only pursues a greener catalytic approach but also allows to target bionano science. Thanks to the multiple mechanisms of molecular amplification (e.g., enzyme turnover, radical chain propagations) and the capability to localize the polymerization at microscale level, new applications for biosensing, high-throughput screening, and chemical imaging emerged (Malinowska & Nash, 2016). As an example, HRP-mediated polymerization was exploited to prepare molecularly imprinted polymers (MIP) for the recognition of 2,4-dichlorophenoxyacetic acid (an herbicide) and salicylic acid, starting from the immobilized HRP catalyst in combination with H2O2, methacrylate, vinyl monomers, cross-linkers and acetylacetone as the mediator (Daoud Attieh, Zhao, Elkak, Falcimaigne-Cordin, & Haupt, 2017). The capability of HRP to promote the formation of a fluorescent alginate hydrogel in the presence of hydrogen peroxide, produced in situ by glucose oxidase (GOx), was recently used by Nash and coworkers to developed a platform for the directed evolution of GOx (Vanella, Ta, & Nash, 2019). The mechanisms by which these reactions occur are driven by the heme prosthetic group found in the enzymes’ active site (Fig. 1). Exogenous hydrogen peroxide (H2O2) is converted into a usable radical source especially when the reaction is coupled with β-diketone mediators. These polymerizations proceed through the mechanism by which nature processes peroxides, the oxidoreductive pathway. Utilizing hydrogen peroxide, a hydroxyl radical is generated which reacts with the mediator acetylacetone to a more stable acetylacetone-based radical. The β-diketone mediator plays two roles in the reaction: (1) It is a reducing substrate for the enzyme, and (2) it forms a radical capable of initiating the polymerization of vinyl monomers (Durand, Lalot, Brigodiot, & Marechal, 2000; Emery, Lalot, Brigodiot, & Marechal, 1997). This radical species then initiates the chain growth process. HRP undergoes its biological role when exposed to peroxides, forming compound I and compound II before it returns to its original FeIII species which is required for peroxidase activity. Furthermore, compared to AIBN or photo initiated FRPs, HRP-initiated polymerizations offer access to the same materials but under benign reaction conditions without the need for elevated temperatures or photo irradiation (Hollmann, 2010; Hollmann & Arends, 2012; Kobayashi, Uyama, & Kimura, 2001). This can be highly advantageous for the production of temperature and light sensitive materials.
Fig. 1 Catalytic cycle for the free radical polymerization of vinyl monomers using HRP as a catalyst. (M, monomer; Pn, propagating polymer radical; kp, rate constant of propagation (i.e., polymer chain growth)).
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Herein, we describe a comprehensive synthetic strategy for enzymeinitiated free radical polymerizations of vinyl monomers that is based on work by Marechal (Durand et al., 2000; Emery et al., 1997; Lalot, Brigodiot, & Marechal, 1999), Gross (Kalra & Gross, 2000) and Kobayashi (Kobayashi et al., 2001). The aim is to provide the reader with know-how on what is required for generating polymeric materials using this method.
2. Free radical polymerizations of vinyl monomers using horseradish peroxidase 2.1 Polymerizations Equipment • Analytical balance • Schlenk line outfitted with five ports equipped with inert gas (Ar or N2) • Schlenk flask (10 mL) • Stirring plate • Magnetic stirring bar • Syringe (1 and 2 mL) • Micropipette (20, 100, 1000 μL) • Needles (ø 0.80 50 mm and ø 0.80 120 mm) • Glass Pasteur pipette (230 mm) • Cotton or glass wool • Scintillation vial (6 mL) • 3 rubber septum Chemicals • Horseradish peroxidase, (HRP; highly stabilized, essential salt-free, lyophilized powder, 200–300 units/mg solid, P2088) (Sigma) • 4-Acryloylmorpholine (AM) (Sigma) • 2-Hydroxyethyl methacrylate (HEMA) (Sigma) • Poly(ethylene glycol) methyl ether acrylate (PEGA; Mn ¼ 480 g mol1) (Sigma) • Acetylacetone (ACAC) (Sigma) • Hydrogen peroxide (H2O2; 30 wt% in H2O) (Sigma) • Aluminum oxide activated, basic, Brockmann I (Sigma) • Aluminum oxide activated, neutral, Brockmann I (Sigma) • Milli-Q water • Dimethyl sulfoxide (DMSO) (Sigma) • Diethyl ether (Sigma)
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Procedure The free radical polymerizations of vinyl monomers using horseradish peroxidase as a catalyst (Scheme 1) were performed as follows: 1 A Schlenk flask, equipped with a magnetic stirring bar and rubber septum, was attached to a Schlenk line in a fume hood. The flask was then evacuated by vacuum and back filled with argon three times. 2 The monomers were purified of inhibitors by passing them through a small plug of basic aluminum oxide via a positive flow of compressed air into 6 mL scintillation vials. The plugs were prepared by inserting a small piece of cotton into a Pasteur pipette and filling it half way with basic aluminum oxide. 3 To the open Schlenk flask under a slight positive flow of argon, HRP (6.1 mg, 77.6 μmol) was added. 10 mmol monomer (PEGA 5 mmol) was added using a 1000 μL micropipette. Once both were added, 700 μL of Milli-Q water and 300 μL of DMSO were added to the flask to dissolve the HRP and monomer. The flask was then sealed with a rubber septum. 4 Using a ø 0.80 120 mm needle and closing the stopcock of the Schlenk flask, argon was passed through the solution to degas the mixture for 15 min with an exhaust needle (ø 0.80 50 mm) through the septum. 5 In a scintillation vial (6 mL), ACAC (25.0 mg, 0.25 mmol) and H2O2 (8.5 mg, 0.25 mmol) were added using a 100 μL micropipette and 20 μL micropipette, respectively, and dissolved in 700 μL of Milli-Q water and 300 μL of DMSO. The vial was then sealed using a rubber septum and was degassed by inserting a needle (ø 0.80 120 mm) into the solution and passing argon through the liquid for 15 min with an exhaust needle (ø 0.80 50 mm) through the septum. 6 To start the reaction, the contents of the scintillation vial were transferred to the Schlenk flask containing the enzyme and monomer using an argon purged syringe (2 mL). 7 Once all of the reactants were added, a 0.3 mL aliquot of the reaction mixture was taken using an argon purged syringe (1 mL). The aliquot was then passed over a plug of neutral aluminum oxide (prepared the same way as step 2) to quench the reaction and remove the HRP. The plug was rinsed with 1 mL of D2O (for p(HEMA) DMSO-d6) into a scintillation vial (6 mL) for NMR and GPC analysis (Section 3). 8 The reaction mixture was stirred at room temperature for a total of 21 h under argon. 9 After the first hour and the final time point, a 0.3 mL aliquot was taken using an argon purged syringe (1 mL) equipped with a needle (ø 0.80 120 mm) from the reaction mixture for NMR and GPC analysis (Section 3). The aliquot was passed through a plug of neutral aluminum oxide as mentioned in Step 7. After the final time point, the reaction was quenched
Scheme 1 Synthesis of p(AM), p(HEMA), and p(PEGA) under free radical polymerization conditions using HRP as a catalyst.
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by opening the reaction flask to air and turning off the flow of argon. The remaining reaction mixture was redissolved in a minimal amount of DMSO and precipitated into cold diethyl ether. The resulting product was filtered and dried under vacuum.
3. Polymer characterization Equipment • NMR spectrometer, e.g., Bruker Ascend 400 with a 5 mm sample head operating at 400.13 MHz • NMR tubes, e.g., Wilmad 535-PP-7 and tube cap • Processing software, e.g., Mestrelab Research: MestReNova (Mnova) • GPC system, e.g., PSS SECurity system equipped with a refractive index (RI) detector • Scintillation vial (6 mL) • PTFE syringe filter 0.2 μm pore size • Syringe (2 mL) • GPC vials Chemicals • Deuterium oxide (D2O) (Cambridge Isotope Labs) • DMSO-d6 (Cambridge Isotope Labs) • Magnesium sulfate (Sigma) • Lithium bromide (Sigma) • Dimethylformamide (DMF) (Fisher Scientific)
3.1 Nuclear magnetic resonance (NMR) spectroscopy NMR measurements are used to characterize a variety of parameters in polymer chemistry. Here we highlight the use of 1H NMR spectroscopy to determine the conversion of the polymerization. The 1H NMR spectra were recorded in D2O (for p(HEMA) DMSO-d6). The content of the scintillation vials of the aliquots diluted with D2O (for p(HEMA) DMSO-d6) (c.f. Section 2.1, Steps 7 and 8) were transferred into NMR tubes. The measurements were collected on a Bruker Ascend 400 spectrometer operating at 400.13 MHz. The spectra were recorded using the following parameters: relaxation delay: 1 s and transients: 32. The resulting spectra were processed using Mestrelab Research: MestReNova v. 11.0.1–17801. All spectra were baseline and phase corrected. Fig. 2 shows the 1H NMR spectrum of the final time point of p(AM) to illustrate the integrated peak regions to determine the conversion of the polymerization.
1371.34
1.00
9.5
9.0
8.5
8.0
7.5
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6.5
6.0
5.5
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1.0
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−1.0
f1 (ppm)
Fig. 2 1H NMR spectra of the final time point for p(AM); integral regions are used to calculate the extent of monomer conversion.
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The NMR data collected for the polymers provides insight into the extent of monomer conversion of this polymerization method. Traditionally, free radical polymerizations result in high conversions and large molecular weight polymers, and even polymeric gels. It is imperative to understand the spectral regions for analyzing these data. In terms of conversion, it is common in the polymerizations of vinyl monomers to compare a vinyl proton of the monomer to a well-known peak in the polymer to determine conversion. Fig. 2 illustrates this analytical practice using p(AM) as an example. The spectral region 7.00–6.40 ppm corresponds to one vinyl proton of the monomer, and the region 1.90–0.75 ppm corresponds to the two backbone protons adjacent to the morpholino-pendent group. By integrating these two regions, conversion can be calculated using Eq. (1). 0
Z
1 poly
B C B C B C #Hpoly B C 100% Z %Conversion ¼ B Z C B poly mono C @ A + #Hpoly #Hmono
(1)
HRP-assisted free radical polymerizations suffer from slow initiation due to the self-quenching mechanism afforded by the biological pathway of peroxidases, where the resulting hydroxyl radical reacts with itself to quench further radical formation (Durand et al., 2000); therefore, to ensure high conversion, the reactions were allowed to stir for 21 h. Final time points, represented here in Fig. 3, were taken after the allotted reaction time. All three spectra show almost full monomer consumption (Table 1) and broadened peaks corresponding to the polymer.
3.2 Gel permeation chromatography (GPC) GPC measurements are performed to determine the number average molecular weight (Mn) and dispersity (Đ). These data help elucidate the molecular weights achieved during a polymerization. GPC was performed on a PSS SECurity GPC system equipped with an RI detector using DMF (0.05 M LiBr) as an eluent. Measurements were performed at 60 °C with a flow rate of 1.0 mL/min. The system was outfitted with two Agilent PolarGel M columns (ID ¼ 7.5 mm, L ¼ 300 mm, particle size ¼ 8 μm) and an Agilent
9.0
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Fig. 3 1H NMR spectra of final time points of HRP-initiated p(AM) (red), p(HEMA) (green), and p(PEGA) (blue) polymerizations.
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Table 1 Final polymer characterization determined by GPC. Mn (kDa) Polymer Conversion (%)a
Dispersity (ᴆ)
p(AM)
99.9
75.7
2.93
p(HEMA)
96.8
128.2
2.96
97.6
324.3
1.91
p(PEGA) a
1
Determined by H NMR spectroscopy.
PolarGel M guard column (particle size ¼ 8 μm). The calibration was performed using poly(ethylene oxide) (PEO) standards. Procedure GPC samples were prepared as follows: 1. Following NMR analysis, the NMR tubes were collected and their content was transferred to individual scintillation vials (6 mL). 2. 2 mL of GPC elution solvent (DMF + 0.05 M LiBr) was added to each vial. 3. To remove residual H2O/D2O, magnesium sulfate was added to each sample and stirred until dry. 4. The slurry-like mixture was then aspirated using a syringe (2 mL) which was then equipped with a PTFE syringe filter (0.2 μm pore size). The solution was injected into a GPC vial. 5. The samples were then placed in the autosampler, and for each measurement 100 μL of the sample was injected onto the column. The GPC measurements of the final polymers illustrate the power of this synthetic technique for generating high molecular weight polymers with typical dispersities seen for FRPs (Table 1). These data represent the size and uniformity of the polymer chains generated, which can dictate the utility of these materials.
4. Conclusions The use of horseradish peroxidase in free radical polymerization reactions represents a bio-initiation for the polymerization of vinyl monomers. This technique harnesses the power of naturally occurring biological processes for synthetic polymer chemistry. The enzyme facilitates an efficient catalysis, which leads to high conversion of monomer. Moreover, it represents an environmentally friendly catalyst. As reported in this chapter, this technique is robust to a variety of functional groups and provides benign
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reaction conditions for generating various polymers. The simplicity of the reaction set up makes this technique an extremely valuable tool for generating high value, functional materials. While this technique has a variety of advantages compared to conventional FRP methods, it still suffers some limitations. Due to the need to extract the enzymes from their natural source, the scalability of these reactions has so far been limited. Efforts continue to be made to circumvent this method’s shortcomings. Additionally, this method has influenced others to utilize enzymes for more advanced radical polymerization techniques such as ATRP and RAFT by harnessing the promiscuity of metalloproteins and other biomolecules (see chapters “Biocatalytic ATRP in solution and on surfaces” by Rodriguez et al. and “Enzyme-initiated reversible additionfragmentation chain transfer (RAFT) polymerization: Precision polymer synthesis via enzymatic catalysis” by Wang and An). These findings have resulted in the exploration of a variety of enzymes’ ability to catalyze or initiate non-native radical polymerizations that have improved reaction times, efficiencies, and materials that can be generated (Rodriguez et al., 2018).
Acknowledgments This work was supported by the Swiss National Science Foundation through grants Nos. PP00P2_144697, PP00P2_172927 and Ambizione program grant 179865, as well as through the Department of Pure and Applied Chemistry of the University of Strathclyde.
References Binder, W. H. (2019). The past 40 years of macromolecular sciences: Reflections on challenges in synthetic polymer and material science. Macromolecular Rapid Communications, 40(1), 1800610. https://doi.org/10.1002/marc.201800610. Blyth, J., & Hofmann, A. W. (1845). Ueber das Styrol und einige seiner Zersetzungsprodukte. Justus Liebigs Annalen der Chemie, 53(3), 289–329. https://doi.org/10.1002/jlac. 18450530302. Braun, D. (2009). Origins and development of initiation of free radical polymerization processes. International Journal of Polymer Science, 2009, 893234. https://doi.org/10.1155/ 2009/893234. Daoud Attieh, M., Zhao, Y., Elkak, A., Falcimaigne-Cordin, A., & Haupt, K. (2017). Enzyme-initiated free-radical polymerization of molecularly imprinted polymer nanogels on a solid phase with an immobilized radical source. Angewandte Chemie, 56(12), 3339–3343. https://doi.org/10.1002/anie.201612667. Derango, R. A., Chiang, L.-C., Dowbenko, R., & Lasch, J. G. (1992). Enzyme-mediated polymerization of acrylic monomers. Biotechnology Techniques, 6(6), 523–526. https:// doi.org/10.1007/BF02447825. Durand, A., Lalot, T., Brigodiot, M., & Marechal, E. (2000). Enzyme-mediated initiation of acrylamide polymerization: Reaction mechanism. Polymer, 41(23), 8183–8192. https:// doi.org/10.1016/S0032-3861(00)00204-4.
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Emery, O., Lalot, T., Brigodiot, M., & Marechal, E. (1997). Free-radical polymerization of acrylamide by horseradish peroxidase-mediated initiation. Journal of Polymer Science Part A: Polymer Chemistry, 35(15), 3331–3333. https://doi.org/10.1002/(SICI)10990518(19971115)35:15<3331::AID-POLA27>3.0.CO;2-B. Flory, P. J. (1936). Molecular size distribution in linear condensation polymers. Journal of the American Chemical Society, 58(10), 1877–1885. https://doi.org/10.1021/ja01301a016. Gomberg, M. (1897). Tetraphenylmethan. Berichte der Deutschen Chemischen Gesellschaft, 30(2), 2043–2047. https://doi.org/10.1002/cber.189703002177. Hollmann, F. (2010). Enzymatic polymerization of vinylpolymers. In K. Loos (Ed.), Biocatalysis in polymer chemistry (pp. 143–505). Weinheim: Wiley-VCH. Hollmann, F., & Arends, I. W. C. E. (2012). Enzyme initiated radical polymerizations. Polymers, 4(1), 759–793. https://doi.org/10.3390/polym4010759. Kalra, B., & Gross, R. A. (2000). Horseradish peroxidase mediated free radical polymerization of methyl methacrylate. Biomacromolecules, 1(3), 501–505. https://doi.org/10.1021/ bm005576v. Kobayashi, S., Uyama, H., & Kimura, S. (2001). Enzymatic polymerization. Chemical Reviews, 101(12), 3793–3818. https://doi.org/10.1021/cr990121l. Lalot, T., Brigodiot, M., & Marechal, E. (1999). A kinetic approach to acrylamide radical polymerization by horse radish peroxidase-mediated initiation. Polymer International, 48(4), 288–292. https://doi.org/10.1002/(sici)1097-0126(199904)48:4<288::Aidpi145>3.0.Co;2-i. Malinowska, K. H., & Nash, M. A. (2016). Enzyme- and affinity biomolecule-mediated polymerization systems for biological signal amplification and cell screening. Current Opinion in Biotechnology, 39, 68–75. https://doi.org/10.1016/j.copbio.2016.01.007. Mark, H. (1964). Macromolecules: Origin, development, significance. Biophysical Journal, 4(1 Pt. 2), 5–7. https://doi.org/10.1016/S0006-3495(64)86923-X. Parravano, G. (1951). Chain reactions induced by enzymic systems. Journal of the American Chemical Society, 73, 183–184. https://doi.org/10.1021/ja01145a063. Rodriguez, K. J., Gajewska, B., Pollard, J., Pellizzoni, M. M., Fodor, C., & Bruns, N. (2018). Repurposing biocatalysts to control radical polymerizations. ACS Macro Letters, 7(9), 1111–1119. https://doi.org/10.1021/acsmacrolett.8b00561. Simon, E. (1839). Ueber den fl€ ussigen Storax (Styrax liquidus). Annalen der Pharmacie, 31(3), 265–277. https://doi.org/10.1002/jlac.18390310306. Singh, A., Ma, D., & Kaplan, D. L. (2000). Enzyme-mediated free radical polymerization of styrene. Biomacromolecules, 1(4), 592–596. https://doi.org/10.1021/bm005537j. € polymerisation. Berichte der deutschen chemischen Gesellschaft (A and Staudinger, H. (1920). Uber B Series), 53(6), 1073–1085. https://doi.org/10.1002/cber.19200530627. Vanella, R., Ta, D. T., & Nash, M. A. (2019). Enzyme-mediated hydrogel encapsulation of single cells for high-throughput screening and directed evolution of oxidoreductases. Biotechnology and Bioengineering, 116(8), 1878–1886. https://doi.org/10.1002/bit.27002. Zavada, S. R., Battsengel, T., & Scott, T. F. (2016). Radical-mediated enzymatic polymerizations. International Journal of Molecular Sciences, 17(2), 195. https://doi.org/10.3390/ ijms17020195.
Enzyme-initiated free radical polymerizations of vinyl monomers using horseradish peroxidase Kyle J. Rodrigueza, Michela M. Pellizzonia, Robert J. Chadwickb, Chao Guob, Nico Brunsa,b,* a
Adolphe Merkle Institute, University of Fribourg, Fribourg, Switzerland Department of Pure and Applied Chemistry, University of Strathclyde, Glasgow, United Kingdom *Corresponding author: e-mail address: [email protected] b
Contents 1. Introduction 2. Free radical polymerizations of vinyl monomers using horseradish peroxidase 2.1 Polymerizations 3. Polymer characterization 3.1 Nuclear magnetic resonance (NMR) spectroscopy 3.2 Gel permeation chromatography (GPC) 4. Conclusions Acknowledgments References
250 253 253 256 256 258 260 261 261
Abstract In this chapter, we highlight the use of horseradish peroxidase (HRP) as a catalyst to initiate free radical polymerizations of vinyl monomers under benign reaction conditions. A variety of vinyl monomers, including 4-acryloylmorpholine (AM), 2-hydroxyethyl methacrylate (HEMA), and poly(ethylene glycol) methyl ether acrylate (PEGA) were polymerized. The enzyme converts exogenous hydrogen peroxide into a usable radical source, which when coupled with a β-diketone, yields a radical that initiates chain growth in the presence of monomers. The resulting polymers were characterized using nuclear magnetic resonance (NMR) spectroscopy and gel permeation chromatography (GPC). By using enzymatic free radical polymerizations, polymers can be generated in a sustainable, environmentally-friendly, and scalable fashion.
Methods in Enzymology, Volume 627 ISSN 0076-6879 https://doi.org/10.1016/bs.mie.2019.08.013
#
2019 Elsevier Inc. All rights reserved.
249
250
Kyle J. Rodriguez et al.
1. Introduction Free radical polymerizations (FRPs) are used every year worldwide to manufacture almost 45% of synthetic polymers such as rubber and plastics using peroxy compounds, redox systems or azo initiators (Binder, 2019; Braun, 2009). From a historical perspective, FRP was first reported by Simon (1839) and later by Blyth and Hofmann (1845) in 1839 and 1845, respectively. They observed the formation of glass like solids starting from a styrene containing natural resin isolated from the wounded bark of Liquidambar orientalis in the presence of light. The concept of polymers and radicals were introduce in the same century (by Berzelius, Berthelot and Wurtz) even if the meaning and nature of these entities remained ambiguous until the reports of Gomberg (1897) and Staudinger (1920) at the beginning of last century. The mechanism of FRP was postulated by Flory (1936) when several synthetic polymers obtained by condensation of formaldehyde with phenol, urea and proteins (e.g., Bakelite, Pollopas and Galalit), as well as cellulose derivatives were already on the market (Mark, 1964). Nowadays, due to their simple reaction conditions, FRPs have been implemented for generating materials such as hydrogels, drug vehicles, and packaging. These applications have resulted in FRPs being implemented as a common synthetic tool for generating highly valuable materials. Additionally, in the last decades, new methodologies have been developed to control radical polymerizations (reversible-deactivation radical polymerization techniques such as nitroxide mediated polymerizations (NMP), atom transfer radical polymerizations (ATRP), and reversible-addition-fragmentation chain transfer (RAFT) polymerizations) but their application on industrial scale is still limited and FRPs remain the most widely used techniques due to the well establish and simple reaction conditions (Binder, 2019). Incorporating biocatalysis into traditional chemical processes allows for a more sustainable approach for generating materials necessary for the food and pharmaceutical industries where biocompatibility is necessary. Harnessing the power of biology can be a very powerful tool, e.g., to repurpose enzymes in perform non-native transformations, which result in efficient catalysis through biological mechanisms. After the first report in 1951 by Parravano, who reported the polymerization of methyl methacrylate in the presence of Xantine oxidase and formaldehyde (Parravano, 1951) many enzymes like oxidases and peroxidases were found to be able to catalyze radical polymerizations of a variety of monomers ranging from phenolic compounds to those containing vinyl
Enzyme-initiated free radical polymerizations
251
groups (Derango, Chiang, Dowbenko, & Lasch, 1992; Hollmann, 2010; Hollmann & Arends, 2012; Singh, Ma, & Kaplan, 2000; Zavada, Battsengel, & Scott, 2016). The use of biocatalysts to initiate and/or mediate polymerizations not only pursues a greener catalytic approach but also allows to target bionano science. Thanks to the multiple mechanisms of molecular amplification (e.g., enzyme turnover, radical chain propagations) and the capability to localize the polymerization at microscale level, new applications for biosensing, high-throughput screening, and chemical imaging emerged (Malinowska & Nash, 2016). As an example, HRP-mediated polymerization was exploited to prepare molecularly imprinted polymers (MIP) for the recognition of 2,4-dichlorophenoxyacetic acid (an herbicide) and salicylic acid, starting from the immobilized HRP catalyst in combination with H2O2, methacrylate, vinyl monomers, cross-linkers and acetylacetone as the mediator (Daoud Attieh, Zhao, Elkak, Falcimaigne-Cordin, & Haupt, 2017). The capability of HRP to promote the formation of a fluorescent alginate hydrogel in the presence of hydrogen peroxide, produced in situ by glucose oxidase (GOx), was recently used by Nash and coworkers to developed a platform for the directed evolution of GOx (Vanella, Ta, & Nash, 2019). The mechanisms by which these reactions occur are driven by the heme prosthetic group found in the enzymes’ active site (Fig. 1). Exogenous hydrogen peroxide (H2O2) is converted into a usable radical source especially when the reaction is coupled with β-diketone mediators. These polymerizations proceed through the mechanism by which nature processes peroxides, the oxidoreductive pathway. Utilizing hydrogen peroxide, a hydroxyl radical is generated which reacts with the mediator acetylacetone to a more stable acetylacetone-based radical. The β-diketone mediator plays two roles in the reaction: (1) It is a reducing substrate for the enzyme, and (2) it forms a radical capable of initiating the polymerization of vinyl monomers (Durand, Lalot, Brigodiot, & Marechal, 2000; Emery, Lalot, Brigodiot, & Marechal, 1997). This radical species then initiates the chain growth process. HRP undergoes its biological role when exposed to peroxides, forming compound I and compound II before it returns to its original FeIII species which is required for peroxidase activity. Furthermore, compared to AIBN or photo initiated FRPs, HRP-initiated polymerizations offer access to the same materials but under benign reaction conditions without the need for elevated temperatures or photo irradiation (Hollmann, 2010; Hollmann & Arends, 2012; Kobayashi, Uyama, & Kimura, 2001). This can be highly advantageous for the production of temperature and light sensitive materials.
Fig. 1 Catalytic cycle for the free radical polymerization of vinyl monomers using HRP as a catalyst. (M, monomer; Pn, propagating polymer radical; kp, rate constant of propagation (i.e., polymer chain growth)).
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Herein, we describe a comprehensive synthetic strategy for enzymeinitiated free radical polymerizations of vinyl monomers that is based on work by Marechal (Durand et al., 2000; Emery et al., 1997; Lalot, Brigodiot, & Marechal, 1999), Gross (Kalra & Gross, 2000) and Kobayashi (Kobayashi et al., 2001). The aim is to provide the reader with know-how on what is required for generating polymeric materials using this method.
2. Free radical polymerizations of vinyl monomers using horseradish peroxidase 2.1 Polymerizations Equipment • Analytical balance • Schlenk line outfitted with five ports equipped with inert gas (Ar or N2) • Schlenk flask (10 mL) • Stirring plate • Magnetic stirring bar • Syringe (1 and 2 mL) • Micropipette (20, 100, 1000 μL) • Needles (ø 0.80 50 mm and ø 0.80 120 mm) • Glass Pasteur pipette (230 mm) • Cotton or glass wool • Scintillation vial (6 mL) • 3 rubber septum Chemicals • Horseradish peroxidase, (HRP; highly stabilized, essential salt-free, lyophilized powder, 200–300 units/mg solid, P2088) (Sigma) • 4-Acryloylmorpholine (AM) (Sigma) • 2-Hydroxyethyl methacrylate (HEMA) (Sigma) • Poly(ethylene glycol) methyl ether acrylate (PEGA; Mn ¼ 480 g mol1) (Sigma) • Acetylacetone (ACAC) (Sigma) • Hydrogen peroxide (H2O2; 30 wt% in H2O) (Sigma) • Aluminum oxide activated, basic, Brockmann I (Sigma) • Aluminum oxide activated, neutral, Brockmann I (Sigma) • Milli-Q water • Dimethyl sulfoxide (DMSO) (Sigma) • Diethyl ether (Sigma)
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Procedure The free radical polymerizations of vinyl monomers using horseradish peroxidase as a catalyst (Scheme 1) were performed as follows: 1 A Schlenk flask, equipped with a magnetic stirring bar and rubber septum, was attached to a Schlenk line in a fume hood. The flask was then evacuated by vacuum and back filled with argon three times. 2 The monomers were purified of inhibitors by passing them through a small plug of basic aluminum oxide via a positive flow of compressed air into 6 mL scintillation vials. The plugs were prepared by inserting a small piece of cotton into a Pasteur pipette and filling it half way with basic aluminum oxide. 3 To the open Schlenk flask under a slight positive flow of argon, HRP (6.1 mg, 77.6 μmol) was added. 10 mmol monomer (PEGA 5 mmol) was added using a 1000 μL micropipette. Once both were added, 700 μL of Milli-Q water and 300 μL of DMSO were added to the flask to dissolve the HRP and monomer. The flask was then sealed with a rubber septum. 4 Using a ø 0.80 120 mm needle and closing the stopcock of the Schlenk flask, argon was passed through the solution to degas the mixture for 15 min with an exhaust needle (ø 0.80 50 mm) through the septum. 5 In a scintillation vial (6 mL), ACAC (25.0 mg, 0.25 mmol) and H2O2 (8.5 mg, 0.25 mmol) were added using a 100 μL micropipette and 20 μL micropipette, respectively, and dissolved in 700 μL of Milli-Q water and 300 μL of DMSO. The vial was then sealed using a rubber septum and was degassed by inserting a needle (ø 0.80 120 mm) into the solution and passing argon through the liquid for 15 min with an exhaust needle (ø 0.80 50 mm) through the septum. 6 To start the reaction, the contents of the scintillation vial were transferred to the Schlenk flask containing the enzyme and monomer using an argon purged syringe (2 mL). 7 Once all of the reactants were added, a 0.3 mL aliquot of the reaction mixture was taken using an argon purged syringe (1 mL). The aliquot was then passed over a plug of neutral aluminum oxide (prepared the same way as step 2) to quench the reaction and remove the HRP. The plug was rinsed with 1 mL of D2O (for p(HEMA) DMSO-d6) into a scintillation vial (6 mL) for NMR and GPC analysis (Section 3). 8 The reaction mixture was stirred at room temperature for a total of 21 h under argon. 9 After the first hour and the final time point, a 0.3 mL aliquot was taken using an argon purged syringe (1 mL) equipped with a needle (ø 0.80 120 mm) from the reaction mixture for NMR and GPC analysis (Section 3). The aliquot was passed through a plug of neutral aluminum oxide as mentioned in Step 7. After the final time point, the reaction was quenched
Scheme 1 Synthesis of p(AM), p(HEMA), and p(PEGA) under free radical polymerization conditions using HRP as a catalyst.
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by opening the reaction flask to air and turning off the flow of argon. The remaining reaction mixture was redissolved in a minimal amount of DMSO and precipitated into cold diethyl ether. The resulting product was filtered and dried under vacuum.
3. Polymer characterization Equipment • NMR spectrometer, e.g., Bruker Ascend 400 with a 5 mm sample head operating at 400.13 MHz • NMR tubes, e.g., Wilmad 535-PP-7 and tube cap • Processing software, e.g., Mestrelab Research: MestReNova (Mnova) • GPC system, e.g., PSS SECurity system equipped with a refractive index (RI) detector • Scintillation vial (6 mL) • PTFE syringe filter 0.2 μm pore size • Syringe (2 mL) • GPC vials Chemicals • Deuterium oxide (D2O) (Cambridge Isotope Labs) • DMSO-d6 (Cambridge Isotope Labs) • Magnesium sulfate (Sigma) • Lithium bromide (Sigma) • Dimethylformamide (DMF) (Fisher Scientific)
3.1 Nuclear magnetic resonance (NMR) spectroscopy NMR measurements are used to characterize a variety of parameters in polymer chemistry. Here we highlight the use of 1H NMR spectroscopy to determine the conversion of the polymerization. The 1H NMR spectra were recorded in D2O (for p(HEMA) DMSO-d6). The content of the scintillation vials of the aliquots diluted with D2O (for p(HEMA) DMSO-d6) (c.f. Section 2.1, Steps 7 and 8) were transferred into NMR tubes. The measurements were collected on a Bruker Ascend 400 spectrometer operating at 400.13 MHz. The spectra were recorded using the following parameters: relaxation delay: 1 s and transients: 32. The resulting spectra were processed using Mestrelab Research: MestReNova v. 11.0.1–17801. All spectra were baseline and phase corrected. Fig. 2 shows the 1H NMR spectrum of the final time point of p(AM) to illustrate the integrated peak regions to determine the conversion of the polymerization.
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Fig. 2 1H NMR spectra of the final time point for p(AM); integral regions are used to calculate the extent of monomer conversion.
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The NMR data collected for the polymers provides insight into the extent of monomer conversion of this polymerization method. Traditionally, free radical polymerizations result in high conversions and large molecular weight polymers, and even polymeric gels. It is imperative to understand the spectral regions for analyzing these data. In terms of conversion, it is common in the polymerizations of vinyl monomers to compare a vinyl proton of the monomer to a well-known peak in the polymer to determine conversion. Fig. 2 illustrates this analytical practice using p(AM) as an example. The spectral region 7.00–6.40 ppm corresponds to one vinyl proton of the monomer, and the region 1.90–0.75 ppm corresponds to the two backbone protons adjacent to the morpholino-pendent group. By integrating these two regions, conversion can be calculated using Eq. (1). 0
Z
1 poly
B C B C B C #Hpoly B C 100% Z %Conversion ¼ B Z C B poly mono C @ A + #Hpoly #Hmono
(1)
HRP-assisted free radical polymerizations suffer from slow initiation due to the self-quenching mechanism afforded by the biological pathway of peroxidases, where the resulting hydroxyl radical reacts with itself to quench further radical formation (Durand et al., 2000); therefore, to ensure high conversion, the reactions were allowed to stir for 21 h. Final time points, represented here in Fig. 3, were taken after the allotted reaction time. All three spectra show almost full monomer consumption (Table 1) and broadened peaks corresponding to the polymer.
3.2 Gel permeation chromatography (GPC) GPC measurements are performed to determine the number average molecular weight (Mn) and dispersity (Đ). These data help elucidate the molecular weights achieved during a polymerization. GPC was performed on a PSS SECurity GPC system equipped with an RI detector using DMF (0.05 M LiBr) as an eluent. Measurements were performed at 60 °C with a flow rate of 1.0 mL/min. The system was outfitted with two Agilent PolarGel M columns (ID ¼ 7.5 mm, L ¼ 300 mm, particle size ¼ 8 μm) and an Agilent
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Fig. 3 1H NMR spectra of final time points of HRP-initiated p(AM) (red), p(HEMA) (green), and p(PEGA) (blue) polymerizations.
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Table 1 Final polymer characterization determined by GPC. Mn (kDa) Polymer Conversion (%)a
Dispersity (ᴆ)
p(AM)
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75.7
2.93
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1
Determined by H NMR spectroscopy.
PolarGel M guard column (particle size ¼ 8 μm). The calibration was performed using poly(ethylene oxide) (PEO) standards. Procedure GPC samples were prepared as follows: 1. Following NMR analysis, the NMR tubes were collected and their content was transferred to individual scintillation vials (6 mL). 2. 2 mL of GPC elution solvent (DMF + 0.05 M LiBr) was added to each vial. 3. To remove residual H2O/D2O, magnesium sulfate was added to each sample and stirred until dry. 4. The slurry-like mixture was then aspirated using a syringe (2 mL) which was then equipped with a PTFE syringe filter (0.2 μm pore size). The solution was injected into a GPC vial. 5. The samples were then placed in the autosampler, and for each measurement 100 μL of the sample was injected onto the column. The GPC measurements of the final polymers illustrate the power of this synthetic technique for generating high molecular weight polymers with typical dispersities seen for FRPs (Table 1). These data represent the size and uniformity of the polymer chains generated, which can dictate the utility of these materials.
4. Conclusions The use of horseradish peroxidase in free radical polymerization reactions represents a bio-initiation for the polymerization of vinyl monomers. This technique harnesses the power of naturally occurring biological processes for synthetic polymer chemistry. The enzyme facilitates an efficient catalysis, which leads to high conversion of monomer. Moreover, it represents an environmentally friendly catalyst. As reported in this chapter, this technique is robust to a variety of functional groups and provides benign
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reaction conditions for generating various polymers. The simplicity of the reaction set up makes this technique an extremely valuable tool for generating high value, functional materials. While this technique has a variety of advantages compared to conventional FRP methods, it still suffers some limitations. Due to the need to extract the enzymes from their natural source, the scalability of these reactions has so far been limited. Efforts continue to be made to circumvent this method’s shortcomings. Additionally, this method has influenced others to utilize enzymes for more advanced radical polymerization techniques such as ATRP and RAFT by harnessing the promiscuity of metalloproteins and other biomolecules (see chapters “Biocatalytic ATRP in solution and on surfaces” by Rodriguez et al. and “Enzyme-initiated reversible additionfragmentation chain transfer (RAFT) polymerization: Precision polymer synthesis via enzymatic catalysis” by Wang and An). These findings have resulted in the exploration of a variety of enzymes’ ability to catalyze or initiate non-native radical polymerizations that have improved reaction times, efficiencies, and materials that can be generated (Rodriguez et al., 2018).
Acknowledgments This work was supported by the Swiss National Science Foundation through grants Nos. PP00P2_144697, PP00P2_172927 and Ambizione program grant 179865, as well as through the Department of Pure and Applied Chemistry of the University of Strathclyde.
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