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Confocal Raman Microscopy for the Determination of Protein and Quaternary Ammonium Ion Loadings in Biocatalytic Membranes for Electrochemical Energy Conversion and Storage Rong Cai, Sofiene Abdellaoui, Jay P. Kitt, Cullen Irvine, Joel Harris, Shelley D. Minteer, and Carol Korzeniewski Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.7b03380 • Publication Date (Web): 14 Nov 2017 Downloaded from http://pubs.acs.org on November 15, 2017

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Analytical Chemistry

Confocal Raman Microscopy for the Determination of Protein and Quaternary Ammonium Ion Loadings in Biocatalytic Membranes for Electrochemical Energy Conversion and Storage

Rong Caia, Sofiene Abdellaouia, Jay P. Kitta, Cullen Irvinea, Joel M. Harrisa, Shelley D. Minteera*, and Carol Korzeniewskia,b* a

Department of Chemistry, University of Utah, 315 S 1400, Salt Lake City, UT 84112

b

Department of Chemistry, Texas Tech University, Lubbock, TX 79416

Corresponding Author Email: [email protected] and [email protected] ABSTRACT: The need to immobilize active enzyme, while ensuring high rates of substrate turnover and electronic charge transfer with an electrode, is a centrally important challenge in the field of bioelectrocatalysis. In this work, we demonstrate the use of confocal Raman microscopy as a tool for quantitation and molecular-scale structural characterization of ionomers and proteins within biocatalytic membranes to aid in the development of energy efficient biofuel cells. A set of recently available short side chain Aquivion ionomers spanning a range of equivalent weight (EW) suitable for enzyme immobilization was investigated. Aquivion ionomers (790 EW, 830 EW and 980 EW) received in the proton-exchanged (SO3H) form were treated with tetra-nbutylammonium bromide (TBAB) to neutralize the ionomer and expand the size of ionic domains for enzyme incorporation. Through the use of confocal Raman microscopy, membrane TBA+ ion content was predicted in calibration studies to within a few percent of the conventional titrimetric method across the full range of TBA+ : SO3- ratios of practical interest (0.1 to 1.7). Protein incorporation into membranes was quantified at the levels expected in biofuel cell electrodes. Furthermore, features associated with the catalytically active, enzyme-coordinated copper center were evident between 400 cm-1 - 500 cm-1 in spectra of laccase catalytic membranes, demonstrating the potential to interrogate mechanistic chemistry at the enzyme active site of biocathodes under fuel cell reaction conditions. When benchmarked against the 1100 EW Nafion ionomer in glucose/air enzymatic fuel cells (EFCs), EFCs with laccase airbreathing cathodes prepared from TBA+ modified Aquivion ionomers were able to reach maximum power densities (Pmax) up to 1.5 times higher than EFCs constructed with cathodes prepared from TBA+ modified Nafion. The improved performance of EFCs containing the short

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side chain Aquivion ionomers relative to Nafion is traced to effects of ionomer ion-exchange capacity (IEC, where IEC = EW-1), where the greater density of SO3- moieties in the Aquivion materials produces an environment more favourable to mass transport and higher TBA+ concentrations.

Introduction Confocal Raman microscopy is a versatile technique that provides insights into the structure and composition of materials with a spatial resolution approaching the diffraction limit of the excitation beam.1-5 The aperture of a confocal Raman microscope defines a fixed volume within a sample, usually not more than a few femtoliters (fL)3,6,7, from which Raman scattering is detected. Collection of the scattered radiation and analysis of the resulting confocal Raman spectrum allows the sample composition within the well-defined probe volume to be determined.1,2,6-16 Confocal Raman microscopy has been adapted for spatial profiling of thin (∼200 µm) polymer membranes1,2 and individual (∼75 µm) polystyrene beads.8 The ability to quantify molecules within individual porous silica particles (∼10 µm)6,10,11 and phospholipid vesicles (∼1 µm)14,15,17 has led to sensitive and rapid detection of low molecular weight compounds (i.e., drugs, trace impurities and contaminants) through extraction and preconcentration of the analyte into the ultra-small (sub-picoliter) particle volumes10,11,14,15,17 Confocal Raman microscopy has also been applied successfully to investigate the structure of polymer electrolyte membrane fuel cells where the structure, state of hydration, and transport properties of the ionomer membranes could be characterized12,13,16 In the present work, we explore the potential of confocal Raman microscopy to aid the development of biocatalytic membrane materials suitable for biofuel cell membrane applications. For many years, research in bioelectrocatalysis has focused on strategies for immobilizing

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enzymes at electrodes in a manner that achieves rapid, efficient transfer of charge between enzyme substrates and the electrode while maintaining enzyme stability. Applications have mainly been in relation to amperometric biosensors and biofuel cells. Of these, amperometric biosensor technology was the first to be commercialized, owing in large part to the disposable nature of the biocatalyst component (e.g., diabetic glucose monitors employ glucose oxidase or glucose dehydrogenase immobilized on a carbon electrode and integrated into a disposable test strip18). Biofuel cells, in contrast, have been more difficult to advance, because long biocatalyst lifetimes are required for most energy conversion applications19 In striving to improve the stability of biofuel cells, the design of polymers used for enzyme immobilization has been an area of interest to many researchers20-29 Perfluorosulfonic acid ionomers (PFSAs) comprise a common class of polymer materials often employed for enzyme immobilization at electrode surfaces30-35 PFSAs consist of a polytetrafluoroethylene (PTFE) backbone and perfluorinated side chains terminating in a sulfonic acid group (Scheme 1). In forming membranes and thin films, the PFSA backbone provides mechanical strength, while the flexible side chains aggregate to form hydrophilic domains that facilitate the transport of water and mobile cations. For several decades, Nafion (Scheme 1a), a so-called long side chain PFSA36, has been considered as a benchmark among fluorinated ionomers. In adapting Nafion for enzyme immobilization, our group has shown that quaternary ammonium bromide salts enable Nafion membrane structure to be manipulated34, enlarging ionic clusters (i.e., micellar pockets and hydrophilic domains) to facilitate enzyme incorporation and enhance transport of substrate and reaction products.34,37,38 The exchange of protons for alkylammonium ions also neutralizes the membrane, and the bulky ions provide resistance to pH change.34,37,38 Structurally similar to Nafion, Aquivion ionomer (Scheme 1b) has

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shorter side chains that alter the phase segregation between hydrophobic and hydrophilic domains.36,39,40 With a simple and low cost synthesis route for the base monomer41, Aquivion shows promise as an alternative to Nafion for enzyme immobilization within biofuel cell electrodes.

Scheme 1. PFSA ionomers Nafion (a) and Aquivion (b). For all materials, n ≈ 1.

The current work reports on the preparation of Aquivion as a matrix for a biocatalytic membrane while using confocal Raman microscopy to investigate the membrane composition and structure. Dispersions in the range of 980-790 EW (m = 7.0 – 5.1, Scheme 1b) were investigated. The enzyme laccase, a benchmark O2 reduction catalyst, and bovine serum albumin (BSA), a well-characterized and widely available protein, were used in evaluating the sensitivity of Raman microscopy toward the protein component of biofuel cell membranes. Ionomers were exchanged by tetra-n-butylammonium ions (TBA+) to promote protein incorporation through enhanced size of ionic domains and amphiphilic interactions.20,42 Confocal Raman microscopy measurements performed on TBA+-exchanged membranes demonstrate the ability to quantify the membrane TBA+ content and investigate its effects on the inner membrane chemical environment. Experimental Section Reagents and Enzymes. Aquivion PFSA ionomers D79-25BS, D83-24BS and D98-25BS were provided by Solvey Specialty Polymers (Bollatte, Italy). The ionomers were received as 3


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aqueous dispersions in the proton-exchanged form with corresponding EW (polymer content) of 790 g/eq (25 wt %), 830 g/eq (24 wt %) and 980 g/eq (25 wt %), respectively. For the 790 EW Aquivion dispersion employed for calibration studies, the ion exchange capacity (IEC) and density were specified as 1.3 meq/g and 1.14 g/mL, respectively. Nafion dispersion (1100 g/eq, 5 wt % in mixed alcohols) and Nafion NRE-212 membrane were from Sigma-Aldrich (St. Louis, MO USA). Bovine serum albumin (BSA) (Sigma-Aldrich) was used as received without further purification. Laccase (E.C.: 1.10.3.2) from Trametes versicolor (TvLc) (Sigma-Aldrich) was purified for confocal Raman studies as described in the Supporting Information (SI, section S.1) and otherwise was used as received. Expressed laccase from Streptomyces coelicolor (ScLc) was prepared as described in Ref. 43. Flavin adenine dinucleotide (FAD)-dependent glucose dehydrogenase (E.C.: 1.1.99.10) from Aspergillus sp. (FAD-GDH) was obtained from Sekisui Diagnostics (Kent, UK). Compounds used and synthetic steps in the construction of laccase airbreathing cathodes (e.g., anthracene-modified multi-walled carbon nanotubes (MWCNTs)) and FAD-GDH bioanodes (e.g., naphthoquinone linear polyethyleneimine redox hydrogel (1,2-NQ4-glycidyl-LPEI or NQ-4-LPEI)) are detailed in the SI (section S.2 – S.4). Buffers were prepared from citric acid monohydrate (Fisher Chemical, Asheville, NC, USA) and sodium phosphate dibasic (Sigma-Aldrich). Other chemicals, including D-glucose, TBA+ bromide (TBAB), sodium chloride and ethanol were obtained from Sigma-Aldrich as reagent grade or better and used as received, unless otherwise mentioned. Aqueous solutions were prepared from 18 MΩ-cm water delivered from a Milli-Q water purification system. Modification of PFSA Ionomers with TBAB. Ionomer dispersions in water were diluted to 5 % (w/v) in ethanol. Dispersions in water/alcohol mixtures were used as received. Then, to 5 mL of each ∼5 % (w/v) suspension, TBAB (10 mg - 200 mg) was added to produce solutions

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with the following nominal TBA+ fraction relative to the ionomer SO3- content: 0.1, 0.2, 0.35, 0.45, 0.85 and 1.7 (790 EW Aquivion) and 3.0 (1100 EW Nafion). The mixtures were vortexed with ceramic mixing beads at 1500 rpm for 70 min until uniform viscous solutions were obtained. Afterward, the mixtures were poured into plastic weigh boats (3 in × 3 in) and dried overnight at room temperature in a low humidity environment. To remove excess salt, the resulting films were first soaked in deionized water (18 MΩ-cm), for at least 24 hr, and subsequently rinsed by flushing with fresh deionized water three times. Finally, the films were dried at room temperature for at least 16 hr and stored at ambient temperature in a dry environment. Dispersions of the TBA+-exchanged ionomer used for electrode construction and Raman characterizations were prepared from the dried films. In clean glass vials, dried TBA+-exchanged ionomer was solubilized in sufficient ethanol to bring the ionomer content in the dispersion to 5 % (w/v). Ionomer dissolution was assisted by vigorous vortexing for approximately 5 min. The dispersions were vortexed again briefly just prior to use. The 790 EW Aquivion dispersions that had been exchanged in the presence of excess TBA+ (1.7 mols TBA+ relative to the ionomer SO3mols) and subsequently rinsed and re-suspended as described above are referred to as “1.7× TBA+ exchanged”. The number of exchange sites available to proton following TBAB modification of ionomer was determined from titration following procedures described previously.44,45 Briefly, from resuspended 5 % (w/v) ionomer in ethanol, films were cast and dried under vacuum. Dried films (ca. 50 mg) were weighed and placed in vials containing 3.0 M NaCl overnight to exchange protons for Na+. The protons exchanged were determined by titration of these solutions with nominally 3.00 x 10-3 M standardized NaOH.

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Protein incorporation into TBA+ exchanged ionomers. Processing followed procedures used in the preparation of enzyme deposition solutions for air-breathing cathodes, omitting anthracene modified MWCNTs (SI, section S.3). Briefly, casting solutions were prepared from 1.7× TBA+ exchanged 790 EW Aquivion ionomer mixed with an equal volume of a citratephosphate buffer solution containing solubilized protein. Vortex / sonication steps were applied to promote thorough mixing. The buffer compositions and protein concentrations used are given in the text and figure legends. Raman Spectroscopy. The confocal Raman microscope has been described in detail previously.10,46,47 Briefly, a Kr+ laser operating at 647.1 nm (Coherent, Santa Clara, CA, USA) was used to excite Raman scattering. The laser beam was expanded using a 4× beam expander (50-25-4X-647, Special Optics Inc., Wharton, NJ), directed into the rear entrance port of a Nikon TE 200 inverted microscope (Nikon TE-200, El Segundo, CA, USA), and reflected off of a dichroic mirror (Semrock, Lake Forest, IL, USA), slightly overfilling the back aperture of a 100×, 1.4 numerical aperture oil immersion objective (Nikon Plan Fluor, El Segundo, CA, USA). The beam was directed through the immersion oil and microscope coverslip to bring the focused laser spot to a position within the sample equidistant from the coverslip and the membrane/air interface. Scattered light from the sample was collected using the same microscope objective, and passed back through the dichroic mirror and a final high pass filter (Semrock, Lake Forest, IL, USA) before being focused onto the entrance slit of a grating monochromator (Bruker 500IS, Preston, Victoria, Australia). The diffraction grating, 600 lines/mm blazed at 750 nm, and 50 µm entrance slit provided a spectral resolution of 2 cm-1. Raman scattered light was detected using a charge-coupled device (CCD) camera (Andor, iDus DU401A, South Windsor, CT, USA). The confocal aperture was defined using the entrance slit of the monochromator in the

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horizontal dimension and by binning three rows of pixels on the CCD camera (78 µm) in the vertical dimension.48 The tightly focused excitation beam (d = 0.6 µm) together with the collection aperture in the image plane define a confocal probe volume within the membranes of ∼1.3 fL where 90% detection efficiency was within a depth along the z-direction of ± 0.6 µm.3 The Raman spectra reflect molecular composition within the confocal probe volume and enable quantitation of immobilized enzymes and exchanged TBA+ ions. Raman spectra were obtained by averaging 20 spectra collected with 30 s exposure periods. The spectra were processed using a custom script executed in Matlab (version 8.6; MathWorks, Natick, MA, USA), which subtracted a dark current offset spectrum and subsequently corrected for instrument response49 by ratioing the data to a white-light reference spectrum10. Prior to calibration studies, all spectra were baseline corrected by subtraction of a fourth-order polynomial fitted to the non-peak containing regions of the spectrum (OriginPro 8.0; OriginLab Corp., Northampton, MA, USA). To enable comparison of spectra collected from separate experiments, spectral intensities were normalized8-12,50 using either the strong TFE vibration near 733 cm-1 as an internal standard or the total optical signal integrated across the wavenumber range investigated (normalization is indicated in figure legends). Responses from membrane TBA+ were calibrated from Raman spectra of Aquivion/TBA+ standards prepared from dispersions in ethanol. A response factor (F)11 was determined from the peak scattering at 733 cm-1 (I733), due to the Aquivion TFE centered vibration, and at 1452 cm-1 (I1452), arising from TBA+ ions. Accordingly,11

= ×

(1)

where is the ratio of mols TBA+ to mols ionomer SO3- groups in the standard. In the determination of F values, dispersions with in the range of 0.2 to 1.7 were utilized. F was

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determined from a least-squares fit of the calibration data. Subsequently, for processed membranes containing unknown TBA+ content, the fraction of TBA+ relative to the SO3- groups ( ) was calculated from: = × ′, where = / derived from

Raman spectra of the processed membranes. Electrochemistry. Electrochemistry measurements were performed with a CHI 660E potentiostat/galvanostat (CH Instruments, Austin, TX USA). An I-cell20 was used for both cyclic voltammetry and fuel cell characterizations (see SI, Section S.5). For enzymatic fuel cell (EFC) measurements, the electrolyte filled compartment of the I-cell held a FAD-GDH bioanode (Figure S1). The cell was operated under galvanostatic control with the laccase-modified graphite felt functioning as the cathode. The glucose fuel solution contained 0.1 M glucose in 0.2 M citrate-phosphate buffer (pH 6.5) and was allowed to mutorotate for at least 12 hours before use. Enzymatic fuel cell performance was evaluated under application of a slow current ramp (0.1 µA s-1). Results and Discussion Figure 1 shows confocal Raman spectra of the Aquivion ionomers that are of interest for biofuel cell electrode construction. A spectrum of PTFE is included to enable vibrations centered on TFE units in the backbone to be easily distinguished from Raman scattering

Figure 1. Confocal Raman spectra of protonexchanged Aquivion ionomer as a function of EW. The spectra were baseline corrected and normalized to the total scattering between 150 cm-1 -1425 cm-1. The arrows indicate the direction of peak intensity change as EW decreases. A spectrum of PTFE is plotted beneath the set of Aquivion ionomer spectra.

derived from side chain functional groups. Notable are the strong bands near 731 cm-1, 383 cm-1 and 290 cm-1, and the three features in the

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1200-1400 cm-1 range, all of which correspond to the PTFE backbone.51,52 For the ionomer samples, these features decrease in intensity as ionomer EW becomes smaller. These trends are consistent with the growing contribution of side chains to the total Raman scattering intensity as EW varies from 980 EW (m = 7.0) to 790 EW (m = 5.1) in the area normalized spectra.50 Indeed, the loss in intensity of bands ascribed to ionomer backbone vibrations occurs in parallel with intensity gain in bands near 1176 cm-1, 1058 cm-1, and 971 cm-1 associated with coupled vibrations of side chain sulfonate and ether groups.12,53-55 The 790 EW Aquivion ionomer had superior performance in biofuel cell measurements (vide infra) and thus was the focus of subsequent Raman measurements. An important step in the preparation of biofuel cell electrodes is the exchange of protons for relatively large, hydrophobic cations that have the potential to expand the Figure 2. Confocal Raman spectra of 790 EW Aquivion ionomer membranes in the proton-exchanged form (black) and after partial neutralization by TBA+ ions (red, bright blue, green, dark blue and purple), as indicated. To enable comparisons, after baseline correction the spectra were normalized to the strong 732 cm-1 peak ascribed to ionomer TFE vibrations. Arrows highlight bands associated with TBA+ vibrations and indicate the change in peak intensity with increasing membrane TBA+ content. The inset plots the Raman scattering in the TBA+ peak at 1452 cm-1 as a function of the as-prepared membrane TBA+ : SO3- ratio, expressed as % TBA+. Spectra were recorded with the confocal probe volume positioned within the central region of the membranes. The included error bar shows the standard error from four replicate measurements.

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polymer matrix and create an environment favorable to the incorporation of enzymes. To achieve high enzyme loadings, ionomer dispersions are often ion-exchanged in solutions containing TBAB.34,37,38 The spectra in Figure 2 show


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evidence of TBA+ incorporation into the Aquivion 790 EW ionomer and the ability to quantify changes in membrane TBA+ content based on Raman scattering originating from the confocal probe volume. Each spectrum in Figure 2 was normalized to the dominant Aquivion CF2 stretching feature at 732 cm-1. Accordingly, peaks with strong intensity variation (indicated by upward pointing arrows) are associated with vibrational modes of TBA+ ions56, while bands that remain fixed as the TBA+ concentration changes can be traced to vibrations of the Aquivion ionomer.12 The TBA+ peaks reflect the TBA+ retained by the membrane following ion-exchange and rinsing (see Experimental Section). The inset plots the scattering intensity in the relatively isolated TBA+ peak at 1452 cm-1 versus the as-prepared membrane TBA+ : SO3- ratio, expressed as % TBA+. An analysis of the samples by the conventional titrimetric method (Table 1) shows TBA+ easily replaces proton in the membranes and, over the range of concentrations in Figure 2, is retained by the ionomer with > 90 % extraction efficiency through the aqueous rinsing step. In Figure 3, Raman scattering in the 1452 cm-1 TBA+ peak is plotted versus the actual membrane TBA+ : SO3- ratio determined from the response factor at 1452 cm-1 derived from confocal Raman spectra of calibration standards. Relative to the inset in Figure 2, the plot in

Table 1. Titrimetric analysis results for TBA+ exchange into 790 EW Aquivion

% TBA exchanged (Nominal)

Available sites per gram ionomer (mmol g-1)

% TBA+ exchanged (Titration)

11

1.13 ± 0.01

11 ± 1

23

1.00 ± 0.037

21 ± 3

45

0.62 ± 0.01

51 ± 0.5

85

0.18 ± 0.02

86 ± 1

170

0.013 ± 0.01

99 ± 1

+

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Figure 3 extends to higher TBA+ concentrations. Following earlier studies of alkylammonium ion extraction into Nafion membranes,45 the extent of TBA+ exchange into Aquivion ionomers was determined for conditions that promote the high levels of TBA+ needed to achieve enzyme loadings Figure 3. Raman scattering in the TBA+ peak at 1452 cm-1 as a function of the expected actual membrane TBA+ : SO3- ratio, expressed as % TBA+, determined from confocal Raman spectra of calibration standards. The point labeled “Aq-790 / 1.7× TBAB” indicates the Raman scattering from the calibration standard containing 1.7× TBAB. The error bar shows the standard error from four replicate measurements. The point labeled “Aq790 / 1.7× TBA+ exchanged (rinsed)” indicates responses for 1.7× TBA+ exchanged 790 EW Aquivion ionomer membranes. For the latter point, the open symbol shows the response extrapolated to the calibration line. The intercept of the dashed line with the x-axis indicates the expected membrane TBA+ concentration. See text for further details.

practical for biofuel cell applications.20,34,57-59 For 5 wt % dispersions of 1100 EW Nafion, optimal enzyme loadings can be achieved when TBA+ exchange is performed in the presence a 3-fold stoichiometric excess of TBAB relative to the ionomer IEC.20,34,45,57-59 The conditions, which are close to the limit of TBAB solubility in the ethanol-rich ion-

exchange medium, for 5 wt % dispersions of the 790 EW Aquivion ionomer correspond to a 1.7fold TBAB stoichiometric excess. Figure 3 includes results for a calibration standard containing 1.7× TBAB (point labeled “Aq-790 / 1.7× TBAB”) and 1.7× TBA+ exchanged 790 EW Aquivion ionomer membranes (labeled “Aq-790 / 1.7× TBA+ exchanged (rinsed)”). The scattering from TBA+ in the calibration standard tracks the linear trend of the partially neutralized membranes. The TBA+ amount detected in the 1.7× TBA+ exchanged membranes is lower, consistent with the expected loss during rinsing of any TBAB in excess of the membrane IEC. Extrapolating the data point for this latter sample to the calibration line in the plot indicates the membranes are

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fully TBA+ exchanged, in agreement with the titrimetric analysis results (Table 1, bottom entry (170 %)), but also appear to retain (∼30 %) excess TBAB through non-specific interactions. The titrimetric analysis, which measures ionomer sites available to proton exchange, is insensitive to excess TBA+ ions. While it is possible the low TBAB solubility in the calibration standards may cause predictions of TBA+ membrane content based on the plot in Figure 3 to be somewhat higher than the actual membrane concentrations, as discussed below, excess TBA+ ions may have an important role in helping to facilitate protein uptake by the ionomer. The high level of TBA+ in 1.7× TBA+ exchanged 790 EW Aquivion ionomer membranes is similar to behavior observed for 1100 EW Nafion, where the same conditions of ionomer wt % and TBA+ concentration led to complete TBA+ exchange before rinsing and ca. 90 % (89 ± 3 %) retention of TBA+ after rinsing.45 To investigate the sensitivity of confocal Raman spectroscopy toward membrane enzyme loading, experiments focused initially on the readily available BSA protein. Spectra of BSA incorporated within a 790 EW Aquivion membrane are shown in

Figure 4. (a) Raman spectra plotted over the 1200 cm-1 – 1800 cm-1 region for 790 EW Aquivion ionomer membrane exchanged by TBA+ after casting from aqueous solution in the presence of BSA at 8.5 mg/mL (green) and 1.3 mg/mL (bright blue), as indicated. To enable comparisons, after baseline correction the spectra were normalized to the strong 732 cm-1 peak ascribed to ionomer TFE vibrations. The casting solution was adjusted to pH 3 just below the pI for BSA. The upper left inset (red) shows Raman features recorded from a BSA saturated droplet of citrate buffer. The lower right insets highlight the response through the 1500-1800 cm-1 region for the sample cast from 1.3 mg/mL BSA (bright blue, factor of 5 y-scale enlargement) and a separately prepared 790 EW Aquivion membrane protonexchanged and cast without BSA present (Aq-H+, factor of 10 y-scale enlargement). (b) A calibration plot for the BSA derived from scattering at 1656 cm-1.

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Figure 4a. The BSA amide I vibration (1656 cm-1)60,61 is well separated from bands of the ionomer and TBA+ ions, which dominate the 1500 cm-1 – 1200 cm-1 range. The inset spectrum of BSA collected in this work (Figure 4a, top) matches previously reported Raman spectra of BSA60, where, in addition to the strong amide I band at 1653 cm-1, bands at 1448 cm-1 and 1341 cm-1 corresponding to low energy deformation and scissoring-type modes from CH2 and CH3 groups in BSA aliphatic side chains are also prominent.60,61 These aliphatic group vibrations are, however, difficult to discern in spectra of the membrane immobilized BSA, due to background scattering from the Aquivion-TBA+ matrix; thus analyses were carried out using only the isolated amide I band. The plot in Figure 4b shows the linearity in the scattering response from the BSA amide I vibration for loadings derived from dispersions containing protein in the range of practical interest (ca. 0.5 mg/mL – 15 mg/mL) for biofuel cell electrodes. The figure also includes a segment of a spectrum from hydrated, proton-exchanged ionomer through the water bending vibration near 1638 cm-1. Comparing the intensity in the water peak to the amide I region in the spectrum of the 1.3 mg/mL BSA loaded Aquivion membrane in Figure 4a indicates the signal-to-noise ratio needed for detection of protein amide I vibrations in biofuel cell cathodes is well above scattering interferences from membrane bound water. Figure 5 shows a confocal Raman spectrum of the TvLc enzyme employed in practical laccase biofuel cell cathodes28,62-65 entrapped within a TBAB modified 790 EW Aquivion ionomer membrane. While the majority of features in Figure 5 are traceable to vibrations of Aquivion (i.e., 732 cm-1, 1049 cm-1, 1200-1400 cm-1) and TBA+ (i.e., 1452 cm-1, 880-910 cm-1), the band arising from TvLc due to the amide I vibrations is apparent at 1673 cm-1. The band is toward higher frequency in the range of protein amide I vibrational modes and likely reflects the high beta-sheet content of the enzyme with the possibility of some randomization in

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conformational structure.61,66-69 To confirm the feature as attributable to laccase, spectral measurements were performed on a membrane prepared using a highly purified recombinant ScLc. The segment of the spectrum encompassing the amide I vibration for the recombinant ScLc is included in Figure 5 as an Figure 5. Raman spectrum for 790 EW Aquivion ionomer membrane exchanged by TBA+ after casting in the presence of laccase (TvLc) (1.8 mg/mL). The left inset enlarges the spectral region containing features associated with vibrations of functional groups near the enzyme active site 70. The right inset (bright blue) labeled “ScLc“ shows the amide I region recorded from a membrane prepared as described above except using a highly purified recombinant laccase (ScLc) sample present at 4 mg/mL.

inset and shows the band with good sensitivity. While the expense of the column purified TvLc made it impractical at this point to investigate a wide range of enzyme loadings, the sensitivity toward the amide I vibrational features were

comparable for the laccase enzymes and BSA protein studied (Figures 4-5). The results are encouraging our plans to target a wider range of protein concentration for TvLc, and other enzymes that have practical importance as biofuel cell catalysts. These experiments will be performed in parallel with independent determinations of protein uptake and enzyme activity. Particularly significant in Figure 5 are features associated with vibrational modes of the enzyme-coordinated copper center at the catalyst active site evident at 415 cm-1, 458 cm-1 and 488 cm-1.70,71 These low-frequency vibrational bands are shown on an expanded scale in the left inset within Figure 5 and in comparison to the featureless response for BSA through the region in Figure S2. There is a possibility of increasing the sensitivity of detecting bands reflective of copper-coordination within the enzyme active site through resonance enhancement of the Raman scattering.70,71 Surprisingly, while resonance enhancement would be expected at the laser

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excitation wavelength employed in these measurements, the bands between 400 cm-1 – 500 cm-1 are of relatively low scattering intensity (Figure 5). This result may reflect some enzyme denaturation, leading to a loss or decrease of the electronic absorption band associated with resonance enhancement. The recombinant ScLc, in fact, had very poor oxygen reduction activity and correspondingly weak bands through the 400 cm-1 – 500 cm-1 region. We anticipate isolating higher activity enzyme at the purification stage may produce stronger resonance-enhanced Raman scattering through this spectral region in measurements on laccase biocatalytic membranes. Although the loading of column purified TvLc had to be limited in membranes prepared for the confocal Raman measurements reported in Figures 5 and S2, the spectral sensitivity was sufficient for detection of key protein vibrations at membrane protein concentrations 5-10 times lower than the enzyme (i.e., laccase and other multi-copper oxidases) levels of a typical airbreathing biofuel cell cathode.58,72 The sensitivity toward the amide I modes, and in particular, the ability to detect vibrations in the vicinity of the enzyme active site have encouraged our next steps in adapting confocal Raman microscopy to the study of practical laccase biofuel cell cathodes. In addition to expected benefits of laccase quantitation, there is potential to uncover mechanistic chemistry occurring at the enzyme active site under reaction conditions typical for energy conversion. Furthermore, the strategies developed will provide a blueprint for approaching other biocathode platforms of interest, such as those based on bilirubin oxidase58,72,73 or nitrogenase.74-76 Returning to Figure 3, the red square indicates the scattering at 1452 cm-1 arising from remaining TBA+ in a 790 EW Aquivion membrane following BSA immobilization. The difference in scattering intensities between the 1.7× TBA+ (rinsed) and the BSA modified

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membranes is proportional to the amount of TBA+ displaced during processing. Confocal Raman spectra of the 1.7× TBA+ and protein loaded membranes over the range of the TBA+ 1452 cm-1 and protein amide I peaks are enlarged in Figure S3 and demonstrate the sensitivity toward changes in membrane TBA+ that accompany biomolecule incorporation. The TBA+ loss reflects its displacement by protein as well as extraction of any excess TBAB into the protein solution buffer and exchange of buffer ions into the ionomer. Given TBA+ (∼1 nm diameter)77 occupies a volume about 250 times smaller than BSA78,79 and TvLc80, then for a constant film volume each protein macromolecule entering the ionomer matrix can be expected to displace several hundred TBA+ ions. The large quantity of TBA+ displaced by protein entering the ionomer at a constant matrix volume suggests the TBA+ stoichiometric excess (ca. 2- to 3-fold for 1100 EW Nafion20,34,57) required during ion-exchange to achieve appreciable protein uptake derives from a need for excess mobile ions (i.e., TBA+, Br- and buffer ions) to minimize the role of protein in matrix charge compensation. With considerable TBA+ availability, protein transport into the ionomer matrix can be driven by extraction into the hydrophobic environment, with excess mobile ions providing charge balance. Laccase air-breathing cathodes were constructed from the TBA+ modified ionomers. In the air-breathing cathode configuration, ionomers form a framework that encapsulates entrapped enzymes and stabilizes linkages81,82 which transfer electrons between the enzyme active sites and the graphite felt current collector (Figure S1). The fluorinated ionomer framework also facilitates the transport of ions and water molecules across the cell and presents a low barrier for the diffusion of O2 into the matrix from the contacting atmosphere. Figure 6 compares the performance of TvLc cathodes prepared from Aquivion ionomers and the 1100 EW Nafion benchmark material within glucose/air EFCs. In these experiments, the air-breathing TvLc

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biocathode was directly exposed to the laboratory atmosphere, while the bioanode side of the electrochemical cell was filled with glucose-containing electrolyte20 (Figure S1). The bioanode consisted of FAD-GDH immobilized within a naphthoquinone-based Figure 6. Representative polarization (left yaxis) and power density (right y-axis) curves for glucose/O2 EFCs employing a TvLc airbreathing biocathode prepared from the indicated ionomers. The EFCs were operated in ambient air with a NQ-4-LPEI GDH bioanode in quiescent buffer (0.2 M citrate-phosphate pH 6.5) containing 0.1 M glucose. Control measurements were performed with denatured laccase.

redox polymer (NQ-LPEI GDH anode) facilitating the mediated bioelectrocatalytic oxidation of glucose at relatively low potential (-0.23 V versus a saturated calomel electrode reference).83 Figure 6 shows representative

polarization and power curves measured by galvanostatic discharge under quiescent conditions. Additionally, a control was performed with denatured TvLc. From these experiments, the maximum current density (Jmax), open circuit potential (OCP) and maximum power density (Pmax) were calculated for each EFC. Table 2 reports the averages of these metrics obtained in triplicate measurements. The variability in OCP for the different ionomers evident in Figure 6 is reflected in the uncertainties (ca. ± 40 mV) reported in Table 2. The results in Figure 6 and Table 2 show that biocathodes prepared with Aquivion ionomers exchanged by TBA+ exhibit higher performances over those constructed from TBA+ modified Nafion. The Pmax values for EFCs prepared from Aquivion dispersions increased with the ionomer IEC, ranging from 63 ± 7 µW cm-2 (790 EW) to 48 ± 5 µW cm-2 (980 EW). The TBA+ exchanged Aquivion ionomers were able to reach Pmax values of up to 1.5 times higher than the Pmax obtained with the similarly prepared Nafion (Pmax = 42 ± 8 µW cm-2). Furthermore, the best

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Table 2. Comparison of glucose / O2 EFC performance metricsa Ionomers Aquivion: 790 EW 830 EW 980 EW Nafion: 1100 EW a

OCP

Jmax

IEC

Pmax -2

(V)

(mA cm )

(µ µW cm )

(mmol g-1)

0.82 ± 0.04 0.83 ± 0.01 0.80 ± 0.04

0.15 ± 0.01 0.14 ± 0.01 0.12 ± 0.01

63 ± 7 61 ± 5 48 ± 5

1.3 1.2 1.0

0.77 ± 0.02

0.11 ± 0.02

42 ± 8

0.9

-2

The values reported are averages and standard errors determined from triplicate measurements.

performance in terms of Jmax and Pmax values were yielded by EFCs prepared with the 790 EW Aquivion (Jmax = 0.15 ± 0.01 mA cm-2; Pmax = 63 ± 7 µW cm-2), which has the greatest density of sulfonic acid moieties among the ionomers examined. The trends in Figure 6 and Table 2 track the ionomer IEC. For the Aquivion ionomers, the IEC scales directly with the frequency of side chains relative to the backbone TFE groups (Figure 1). Referring to Scheme 1, m on average for the 790 EW and 830 EW Aquivion ionomers is 5.1 and 5.5, respectively, whereas, the side chains in the 980 EW Aquivion (and 1100 EW Nafion) have 1-2 TFE units greater separation. Although 1100 EW Nafion and 980 EW Aquivion have similar m, the longer side chains in Nafion result in a lower IEC. The hydrophilic properties and plasticizing effects imparted by the side chain sulfonate and ether groups reduce resistance to mass transport within the ionomer framework and are expected to play an important role affecting the biofuel cell performance metrics.84-86 Similar trends are evident in the catalytic currents for O2 reduction recorded in half-cell voltammetric measurements on laccase cathodes (See SI section S.8). However, in addition to improving mass transport, the sulfonate groups also support high levels of TBA+ uptake, which has the potential to produce larger membrane pores and enzyme loadings. Raman spectral measurements 18



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confirmed the membrane processing conditions for the 790 EW, 830 EW and 980 EW Aquivion ionomers and the 1100 EW Nafion ionomer led to full exchange by TBA+. Laccase quantitation, however, was complicated by background fluorescence from impurities in the protein extract used for biofuel cell and voltammetry studies. Although the fluorescence interference prominent in membranes cast from enzyme deposition solutions onto microscope coverslips did not persist in confocal Raman measurements on fully constructed EFC cathodes, heterogeneity in the protein coverage on the cathode led to large uncertainties in the prediction of TvLc amounts. The question of whether the TBA+ amounts, which scale with ionomer IEC, affect protein loadings is being addressed further as the technique is adapted to in situ measurements on biofuel cell electrodes. Conclusions A confocal Raman microscope equipped with an oil immersion objective demonstrated the sensitivity and probe volume (∼1.3 fL) spatial characteristics needed to support the study of composition and active site structure within biocatalytic fuel cell membranes. When applied in conjunction with electrochemical measurements on newly available Aquivion ionomers, confocal Raman measurements showed the up to 3-fold stoichiometric excess of alkyl ammonium ions (e.g., TBA+)20,34,57 required during processing to achieve high enzyme loadings results in full TBA+ exchange with the likelihood for retention of excess TBA+ salts. TBA+ ions are expected to expand the ionomer membrane pores and create a hydrophobic environment that assists enzyme extraction into the polymer.20,34,57-59 The high TBA+ levels in processed membranes revealed by confocal Raman microscopy likely counter the effects of TBA+ depletion during protein uptake and ensure a source of excess mobile ions that can minimize the role of protein in matrix charge compensation. The more favourable mass transport

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characteristics and greater TBA+ concentrations available to increase biocatalyst loadings can explain the improved performance of laccase air-breathing cathodes prepared from the higher IEC Aquivion ionomers (790 EW, 830 EW and 980 EW) relative to 1100 EW Nafion. The results encourage future investigation of Aquivion ionomers in biofuel cell development, including studies of durability and lifetime. Furthermore, detecting vibrational spectral features associated with enzyme coordination of the catalytically-active copper sites of TvLc immobilized within Aquivion membranes could enable studies of laccase active site structure under biofuel cell operating conditions; this measurement approach is suited to other enzymes of interest for biocathode development (i.e., bilirubin oxidase58,72,73 and nitrogenase74-76,87). Nitrogenase is of far-ranging interest as a biocathode catalyst for its ability to transform atmospheric N2.74-76,87 For laccase, bilirubin oxidase and nitrogenase, there is potential to resonantly enhance Raman scattering from moieties near the catalytic centers,70,71,88-91 providing the sensitivity needed to investigate the influence of biofuel cell membrane environment on enzyme active site structure. The reported studies demonstrate confocal Raman microscopy as a valuable technique for guiding advances in biofuel cell membrane processing and design. With near diffraction limited spatial resolution (ca. 0.6 µm in-plane and 1.2 µm depth resolution), in future applications there is potential for quantitative mapping of membrane structure and composition to identify spatial heterogeneity that affects electron and mass transport pathways key to energy conversion efficiency in EFCs. Associated Content Supporting Information Purification of laccase (E.C.: 1.10.3.2) from Trametes versicolor (TvLc); Synthesis of anthracene-modified MWCNTs; Enzymatic air-breathing cathode assembly; FAD-GDH Bioanode; Electrochemical cell used for voltammetry and biofuel cell characterizations;

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Confocal Raman spectra comparing TvLc and BSA features near 400 cm-1; Confocal Raman spectra showing membrane TBA+ displaced by protein; Laccase air-breathing cathode O2 reduction voltammetry; References. Acknowledgments The authors would like to thank the United States Department of Agriculture NIFA grant program and the U.S. Department of Energy (DE-FG03-93ER14333) for partial funding. This work made use of University of Utah shared facilities of the Micron Technology Foundation Inc. Microscopy Suite sponsored by the College of Engineering, Health Sciences Center, Office of the Vice President for Research, and the Utah Science Technology and Research (USTAR) initiative of the State of Utah. This work also made use of University of Utah USTAR shared facilities supported, in part, by the MRSEC Program of the NSF under Award No. DMR1121252. The NSF-DMR award additionally provided REU funding for Mr. Cullen Irvine. C.K. gratefully acknowledges support from the Texas Tech University faculty development leave program. References 1. Everall, N. J. Appl. Spectrosc. 1998, 52, 1498-1504. 2. Everall, N. J. Appl. Spectrosc. 2000, 54, 773-782. 3. Bridges, T.; Houlne, M.; Harris, J. M. Anal. Chem. 2004, 76, 576-584. 4. Opilik, L.; Schmid, T.; Zenobi, R. Annu. Rev. Anal. Chem. 2013, 6, 379-398. 5. Stewart, S.; Priore, R. J.; Nelson, M. P.; Treado, P. J. Annu. Rev. Anal. Chem. 2012, 5, 337360. 6. Bryce, D. A.; Kitt, J. P.; Harris, J. M. Anal. Chem. 2017, 89, 2755-2763. 7. Kitt, J. P.; Bryce, D. A.; Minteer, S. D.; Harris, J. M. J. Am. Chem. Soc. 2017, 139, 38513860. 8. Bridges, T. E.; Uibel, R. H.; Harris, J. M. Anal. Chem. 2006, 78, 2121-2129. 9. Schaefer, J. J.; Ma, C.; Harris, J. M. Anal. Chem. 2012, 84, 9505-9512. 21


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