The work of K.A.V. and P.A.A. was supported by the European Research Council (EnergyBioCatalysis-ERC-2010-StG-258600), Engineering and Physical Sciences Research Council IB Catalyst award EP/N013514/1, and Biotechnology and Biological Sciences Research Council (BB/L009722/1 and BB/N006321/1). R.H. was supported by Ministerio de Ciencia y Tecnolog&#236;a, Universidad de Costa Rica, and Lincoln College, Oxford. The authors acknowledge Mr. Charlie Jones, Mr. Charlie Evans and staff of the Mechanical Workshop (Department of Chemistry) for assistance in the design and manufacture of spectroelectrochemical cells used in this work.

1. Recreating the Internal Sample Compartment of an FTIR Spectrometer Inside an Anaerobic, Dry Glovebox and General Experimental Requirements
<ol>
	<li>Use a commercial FTIR spectrometer equipped with an external mercury cadmium telluride (MCT) detector and ATR accessory. Purge the body of the spectrometer with dry air or dinitrogen, or alternatively use a spectrometer with an evacuated optical bench.</li>
	<li>Place the spectrometer onto a stable, vibration-free table at the same height as an anaerobic (&#60;1 ppm O2), dry (dew point &#60; -75 &#176;C) glovebox. Divert the IR beam of the spectrometer into the glovebox, through an IR transparent window that is large enough to accommodate the defocused beam. 
	NOTE: It is important that the space between the glovebox and spectrometer is also purged or evacuated, especially if a hygroscopic window material, such as NaCl or KBr for example, is used.</li>
	<li>Replicate the internal sample compartment of the spectrometer inside the glovebox.
	<ol>
		<li>Direct the beam inside the glovebox onto an off-axis ellipsoidal mirror, ideally with the same focal length as the internal focusing mirror of the spectrometer. 
		NOTE: It is useful to have two additional plane mirrors placed before this focusing mirror, in order to allow both the height and the direction of the IR beam to be adjusted inside the glovebox; this will offset any inaccuracy in initial placement of the spectrometer.</li>
		<li>Place an external MCT detector, complete with a suitable focusing optic (such as a short focal length ZnSe lens, or off-axis parabolic mirror, with a short focal length) inside the glovebox, positioned such as to replicate the dimensions of the internal sample compartment of the spectrometer.</li>
		<li>Adjust the &#39;focus&#39; of the IR beam to be approximately equidistant between the focusing mirror and the MCT detector.</li>
	</ol>
	</li>
	<li>Cool the dewar of the external MCT detector with liquid nitrogen.</li>
	<li>Mount the ATR accessory into the glovebox sample compartment. Align the input and output focusing optics and ATR accessory to achieve maximum throughput to the MCT detector. If necessary, use an aperture or other suitable attenuation (such as a wire grid) to prevent oversaturation of the detector signal. 
	NOTE: For the data presented here, a five-reflection ATR accessory with all reflective optics and a removable trapezoidal Si internal reflection element (IRE) is used (Crystal GmbH, dimensions ca 5 &#215; 8 &#215; 1 mm3, face angle 39.5&#176;). The IRE is mounted in a removable baseplate machined from polyether ether ketone (PEEK).</li>
	<li>Use a glovebox to house the recreated sample compartment with suitable feedthroughs to allow connection to an externally-housed potentiostat (gas tight BNC connections are useful for this purpose), gas access, and cable(s) to transfer the signal from the MCT detector. It is important to retain any shielding on the detector cable to avoid introducing noise or interference to the measurements.</li>
	<li>Place a peristaltic pump, capable of flow rates greater than 60 mL/min inside the glovebox. A schematic view of the spectrometer, ATR accessory, peristaltic pump and potentiostat setup is shown in Figure 1.</li>
	<li>Construct a spectroelectrochemical cell, such as that shown schematically in Figure 2, that seals onto the ATR accessory. The cell should contain a high surface area counter electrode (Pt wire or gauze), a connection for the working electrode (carbon rod) and a miniature reference electrode. The cell should contain at least two further ports as inlet and outlet for rapid flow of solutions through the cell. 
	NOTE: A miniature saturated calomel reference electrode is ideal for this purpose.36</li>
</ol>
2. Preparation of Carbon Black Particles Modified with E. coli Hydrogenase 1
<ol>
	<li>In a &#39;wet&#39; anaerobic glovebox, mix 20 mg of high surface area carbon black particles (&#62;1,000 m3/g) with 1 mL ultrahigh purity water (resistivity &#62;18 M&#937; cm) in a microcentrifuge tube. Disperse the particles by low-power sonication (&#60;100 W) for at least 15 min, or until the dispersion is uniform and does not sediment within 1 h, to give a carbon black dispersion with a loading of approximately 20 mg/mL.</li>
	<li>Take an aliquot of E. coli hydrogenase 1 (Hyd1, approximately 50 &#181;L, prepared according to a published procedure55) at a concentration of ~7 mg protein/mL and exchange it into a low ionic strength buffer at a pH close to the isoelectric point (for example potassium phosphate, 15 mM, pH 5.8, no additional salt). To achieve best results, use an enzyme preparation that is as active as possible. Perform the buffer exchange by concentration and dilution, in a centrifugal filter device with an appropriate molecular weight cutoff (50 kDa works well for for Hyd1).
	<ol>
		<li>Add the Hyd1 to the filter device.</li>
		<li>Dilute with approximately 450 &#181;L of exchange buffer.</li>
		<li>Reconcentrate to a volume of 50 &#181;L using mild centrifugation (&#60; ~2700 &#215; g) to prevent irreversible precipitation.</li>
		<li>Repeat steps 2.2.2 and 2.2.3 until buffer exchange is complete (typically within ~5 cycles).</li>
	</ol>
	</li>
	<li>Mix a 5 &#181;L volume of 20 mg/mL carbon black dispersion (prepared in 2.1) with the 50 &#181;L aliquot of buffer-exchanged Hyd1. Store the enzyme-particle mixture in a refrigerator (at 0 &#176;C) overnight to allow the enzyme to adsorb. It may be necessary to shake the mixture occasionally to ensure the particles remain dispersed.</li>
	<li>Wash the Hyd1-modified particles with low ionic strength buffer (potassium phosphate, 15 mM, pH 5.8, no additional salt) by successive sedimentation and re-dispersion cycles in a microcentrifuge (~2,700 &#215; g). Repeat this process 3-5 times to remove non-adsorbed enzyme. 
	NOTE: Before the first wash the supernatant should be virtually colorless, indicating good adsorption of enzyme.</li>
	<li>Concentrate the particles to a final volume of ~5 &#181;L, to give a final loading of ~20 mg/mL of enzyme-modified particles. 
	NOTE: The Hyd1-modified particles can be stored at 4 &#176;C for up to two weeks if they remain hydrated; this is best achieved by storing particles diluted with ~50 &#181;L of ultrahigh purity water.</li>
</ol>
3. Preparation for PFIRE Measurements on E. coli Hydrogenase 1
<ol>
	<li>Clean the Si IRE by low-powered sonication (&#60; 100 W), first in H2SO4 (&#60; 90 % w/w) for ~15 min, and then in HNO3 (70 % w/w) for up to 1 hr. This cleaning procedure should lead to a hydrophilic IRE surface. If further cleaning is required the IRE can be placed in piranha solution (a 1:3 ratio of H2O2:H2SO4) to oxidize any remaining organic material. 
	NOTE: Harsh acidic cleaning methods may not be suitable for use with all IRE materials. Piranha solution is highly corrosive and a strong oxidant, surfaces should be reasonably clean before use of piranha solution and care should be taken during preparation and use of piranha solution due to the exothermic nature of the process - always add H2O2 slowly to H2SO4 and refer to the appropriate safety procedures for preparation, use and disposal.</li>
	<li>Rinse the clean IRE in ultrahigh purity water, and dry under a stream of dry nitrogen gas. Carry out all prism handling using clean tweezers to avoid contamination.</li>
	<li>Seal the IRE into the ATR accessory baseplate using a thin strip of electrical-grade silicone sealant. Take care to restrict the sealant to the edges of the IRE. Allow the sealant to dry fully.</li>
	<li>Transfer the ATR accessory baseplate to the spectrometer glovebox and mount it onto the ATR accessory. Measure a reference (background) spectrum between 4000 - 1,000 cm-1 at 4 cm-1 resolution, using the standard rapid scan mode of the spectrometer and 1024 averaged interferograms (measurement time ~3 - 5 min). Use this spectrum as an input to allow computation of absorbance spectra later in the experiment. 
	NOTE: Due to the small absorbance expected for the Hyd1 active site it is necessary to collect spectra with high signal to noise ratios.</li>
	<li>Transfer the ATR accessory baseplate to the &#39;wet&#39; glovebox which contains the pre-prepared Hyd1-modified particle dispersion (section 2). Drop-cast a 1 &#181;L aliquot of the particle dispersion onto the large face of the IRE, and spread them evenly across the surface. Note: Do not allow the particles to become completely dry on the IRE.</li>
	<li>Cut a piece of carbon paper, such as that used as gas diffusion layer material in fuel cells, to a size ~0.1 mm smaller than the surface dimensions of the IRE (i.e. ca. 8.2 &#215; 4.9 mm2). The surface area of the carbon paper should be large enough to cover the drop-cast Hyd1-modified particles, but not so large as to overlap the silicone sealant used to secure the IRE. Soak the carbon paper in water, and gently place it over the top of the particle film. The carbon paper will provide a good electrical connection across the whole particle film. 
	NOTE: It is beneficial to prepare several pieces of carbon paper in advance, and store them pre-soaked in ultrahigh purity water inside the &#39;wet&#39; anaerobic glovebox.</li>
	<li>Mount the spectroelectrochemical cell (described in 1.7 and shown schematically in Figure 2) over the IRE, secure it into the baseplate with screws. Add ~200 &#181;L of the experimental buffer to keep the enzyme hydrated. A mixed buffer system, capable of buffering over a wide pH range, is useful for studies on NiFe hydrogenases:56 sodium acetate, 2-[N&#39;-morpholino]ethane-sulfonic acid (MES), N&#39;-[2-hydroxyethyl]piperazine-N&#39;-[2-ethane-sulfonic acid] (HEPES), N&#39;-tris[hydroxymethyl]methyl-3-amino-propane-sulfonic acid (TAPS), and 2-[N&#39;-cyclohexylamino]ethane-sulfonic acid (CHES), with each component at a final concentration of 15 mM and containing 0.1 M NaCl as supporting electrolyte, with pH adjusted to pH 6 using concentrated NaOH and HCl. 
	NOTE: The working electrode connection should protrude slightly below the plane of the spectroelectrochemical cell top (~0.1 mm) to ensure good electronic connection to the carbon paper (Figure 2).</li>
	<li>Connect the solution inlet and outlet of the spectroelectrochemical cell to a flow system containing peristaltic pump tubing and a vial of the experimental buffer. Transfer the assembled cell to the &#39;dry&#39; glovebox containing the spectrometer.</li>
	<li>Mount the spectroelectrochemical cell assembly onto the ATR accessory. Connect the working, counter and reference electrodes to the potentiostat. Connect the peristaltic pump tubing to the pump.</li>
	<li>Record an absorbance spectrum with 1024 averaged interferograms at 4 cm-1 resolution over a spectral range of 4,000 - 1,000 cm-1, using the spectrum collected in 3.4 as a reference/background. At this point the spectrum should contain significant (&#62;100 mO.D.) amide II bands at ~1,540 cm-1 and active site bands of Hyd1 should be evident in the spectral region 1,850 - 2,150 cm-1, largely in oxidized, inactive states (Figure 3).</li>
</ol>
4. Activation of E. coli Hydrogenase 1 and Testing the Spectroelectrochemical Cell
<ol>
	<li>Apply a reducing potential (&#8722;0.8 V vs SCE) to the Hyd1-modified particle film. Saturate the experimental buffer with H2 and flow buffer slowly through the spectroelectrochemical cell. Leave the sample overnight to fully activate the Hyd1. 
	NOTE: It is important to use anaerobic H2, N2 etc., and so all anaerobic gases should be passed through an O2 filter.</li>
	<li>Record an absorbance spectrum of the sample after activation. The &#957;CO and vCN bands of the active site should now show a distribution of reduced, &#39;active&#39; states. This is most easily observed through the use of a difference spectrum, relative to the spectrum recorded in 3.10 (Figure 4).</li>
	<li>Test the electrical connection of the spectroelectrochemical cell. To do this, saturate the experimental buffer with N2 gas. Apply a sequence of oxidizing (0 V vs SCE) and reducing (&#8722;0.8 V vs SCE) potentials (of ca 30 min duration) to the Hyd1-modified particle film, and record an absorbance spectrum at each. The Hyd1 should become rapidly oxidized and reduced, and if the particle film is well connected 100 % of the sample should respond to the applied potential.</li>
	<li>Set an appropriate flow rate of the experimental buffer. To do this, apply a reducing potential (&#8722;0.8 V vs SCE) to the sample, and saturate the experimental buffer with H2. Record a series of cyclic voltammograms between -0.707 - 0.039 V vs SCE at a scan rate of 10 mV/s. Gradually increase the flow rate of H2-saturated buffer between voltammograms until the catalytic waveshape resembles that at a planar rotating disc electrode55 and the maximum current is independent of flow rate (Figure 5). 
	NOTE: The solubility limit of H2 in water is ~0.8 mM at 293 K and 1 bar.</li>
	<li>As the sample is now ready for PFIRE measurements, collect spectra at a range of potentials, under a range of solution conditions (pH, temperature, H2 concentration etc.). Record all electrochemical data using the potentiostat software as it is important to be able to correlate spectroscopic and electrochemical data, especially when studying electrocatalytic processes such as H2 oxidation by Hyd1.</li>
</ol>
5. Spectroscopic Data Handling
<ol>
	<li>Confirm that the active site has not been permanently altered during the course of the measurements by recording spectra at 0 V and &#8722;0.8 V vs SCE at the end of the experiment. These should be identical to the spectra recorded in 4.3, and no loss of active site should be observed during the measurement (Figure 6).</li>
	<li>Export absolute absorbance spectra from the spectrometer software in a suitable format (.csv, ASCII, &#39;matlab&#39;, Jcamp etc.) for processing using software such as Origin or Matlab.</li>
	<li>Baseline correct the data, using the process illustrated in Figure 7.
	<ol>
		<li>Take the second derivative of each absorbance spectrum in the range 1,800 - 2,150 cm-1, in order to identify small (&#60;1 mO.D.) active site bands against the highly-curved water background.</li>
		<li>Place baseline marker points on the original absorbance spectrum, and set the points to &#39;snap&#39; to the experimental spectrum.</li>
		<li>Fit a baseline through the points using an interpolated cubic spline function or a polynomial function. Subtract this baseline function from the experimental data.</li>
	</ol>
	</li>
</ol>

The authors declare no competing financial interest.

PFIRE is a broadly applicable IR spectroscopic technique for addressing electrode-immobilized redox proteins. In particular, electrocatalytic reactions of redox enzymes can be probed under fast turnover conditions. The PFIRE method builds upon the direct electrochemical control provided by the technique of PFE, which provides no direct structural information, and couples it to IR spectroscopy at a carbon electrode. The PFIRE approach thus adds chemical insight to the information available from electrochemistry alone and is highly suited to study of redox proteins and enzymes involved in small molecule binding and activation. In addition, PFIRE can provide information about potential-dependent structural changes in proteins in the absence of catalytic turnover. Such non-turnover electron transfer events are often difficult to detect using &#39;standard&#39; applications of PFE, although the extension of PFE to Fourier-transformed AC voltammetry has been used with great success.45,46
The PFIRE method is, in principle, suitable for the study of any redox protein that can be studied using PFE. Therefore, as with PFE, protein adsorption is a critical step for a successful PFIRE experiment. In this protocol we describe an application of the PFIRE technique using E. coli Hyd1 as a case study.36,43 However, we have also applied the PFIRE technique to the cytoplasmic regulatory hydrogenase from R. eutropha,44 and to flavin mononucleotide adsorbed on carbon black.40 In all these cases, simple physical adsorption to unmodified high surface area carbon black (as described in this Protocol) provides a surface coverage of protein that is high enough to record good quality IR spectra with a high signal-to-noise ratio. In cases where such high levels of adsorption cannot be achieved it may be necessary to modify the surface of the carbon particles, for example to allow covalent attachment of protein to the electrode surface.60,61,62 The use of a glovebox for PFIRE measurements is only strictly necessary when studying samples that must be handled anaerobically. However in practice the extremely constant and low (&#60; 80 &#176;C dew point) levels of water vapor provided by the glovebox atmosphere give high signal-to-noise levels that allow the extraction of very small absorbances.44 In many cases an anaerobic environment, such as that provided by the glovebox, is also desirable for the electrochemical measurement (integral to the PFIRE technique) in order to avoid current due to O2 reduction at the working electrode.
IR absorbances due to bulk water, experimental buffers and the carbon particles on which the sample is adsorbed all contribute significantly to the experimental spectra and could overlap bands of interest, particularly in the amide I, II and III regions of the spectrum.63 The amide region also contains information from organic species such as flavins or nicotinamide cofactors, as well as the substrates and products of many oxidation and reduction reactions. In the case of NiFe hydrogenases, &#957;CO and &#957;CN bands of the active site fall in a relatively clear region of the spectrum and so the PFIRE technique is very well suited to the study of these enzymes. In other cases, however, difference spectra coupled with isotopic labelling approaches may be needed to isolate changes due to the immobilized protein. Similar approaches have been used to identify, for example, protonation changes, structural rearrangements and to study Michaelis-Menten complexes using IR spectroscopy.64,65,66 PFIRE is not, therefore, limited to the study of hydrogenases but can be applied to any redox protein that contains (or whose substrates, products or inhibitors contain) groups with diagnostic IR-active vibrations; carbon monoxide dehydrogeanses,67 nitrogenases,68,14 flavoproteins,40 and formate dehydrogenases, for example.
The related technique, SEIRA, is very well suited to the study of membrane-associated proteins in a biomimetic environment.32 SEIRA is an adaptation of IR spectroscopy that also uses an ATR-IR configuration, and makes use of a surface enhancement effect that amplifies the IR absorbance of molecules located close to (within a few nm) the surface of the ATR prism (IRE). SEIRA is therefore exquisitely sensitive to spectral changes that occur within the adsorbed protein and membrane architecture and is relatively free from competing signals from solvent and substrates/inhibitors present in solution. This is somewhat in contrast to the PFIRE technique described here which relies on a significantly greater penetration depth above the surface of the IRE (~1 &#181;m), meaning that PFIRE is more sensitive to substrates, products or inhibitors present in solution. This increased sensitivity to &#39;bulk&#39; solvent can be advantageous; if the substrate or product can be observed directly by IR, PFIRE spectra report on both steady state kinetics of long-lived active species and associated product formation during electrocatalysis.69 The ability to observe steady state concentrations of substrate and product will be particularly valuable for enzymes such as carbon monoxide dehydrogenase (which catalyzes the reversible oxidation of CO to CO2, a strong IR absorber) or formate dehydrogenase (which catalyzes the reversible oxidation of formate to CO2).
At present, PFIRE is limited to steady-state kinetic studies of enzyme electrocatalysis due to the macroscopic carbon electrodes used to &#39;concentrate&#39; the enzyme onto the surface of an IRE for data collection in and ATR-IR geometry. In this respect PFIRE studies of NiFe hydrogenases are complementary to the work of Dyer and co-workers,49,50 who use light-triggered transient absorption IR spectroscopy to study sub-turnover kinetics. Work is underway to miniaturize the spectroelectrochemical cell,40 and with the use of microelectrodes time resolution on the order of microseconds should be achievable. This will enable study of sub-turnover kinetics for enzymes with turnover frequencies up to ca 100 - 500 s-1, and will enable the study of both reductive and oxidative processes.
Overall, PFIRE is a spectroscopic technique that allows chemical characterization of electrocatalytic reactions of redox enzymes under steady state conditions. The PFIRE approach allows multiple chemical and electrochemical titrations to be carried out on the same enzyme sample, as the high surface area electrodes used provide a robust adsorption of protein and the ATR-IR geometry allows facile solution exchange. The ability to collect such structural information in situ during enzyme function is an invaluable tool for the wider bioelectrochemistry community.

A key challenge in the study of protein function involves development of in situ methods that allow direct observation of proteins carrying out their physiological roles, either in vivo or using isolated protein samples. This requires the integration of control or triggering processes into experimental procedures and use of combined techniques that allow both the reactivity to be assessed, and individual chemical steps during protein function to be measured, simultaneously. In the case of redox proteins this often equates to combining electrochemical techniques, which precisely control the applied potential but provide no direct chemical information, with spectroscopic techniques that are sensitive to chemically-specific changes associated with protein function.1,2,3 Spectroelectrochemistry is a general term for a range of coupled electrochemical and spectroscopic methods that encompass a variety of spectroscopic techniques and levels of electrochemical control. Many proteins can exchange electrons with artificial, soluble electron donors and acceptors and this has been exploited in studies which use small molecules to mediate electron transfer, including coupling with UV-visible,4,5,6,7 magnetic circular dichroism8 and infrared5,9,10,11,12,13,14 (IR) spectroscopies. In a limited number of cases it has proved possible to exploit unmediated, diffusion-controlled electron exchange between proteins and electrodes.15,16
For catalytic reactions carried out by redox enzymes, solution electrochemistry approaches present a clear disadvantage. Diffusion-controlled electron transfer via redox mediators in solution is likely to become rate limiting. Kinetic and mechanistic information about the enzyme may be lost, or at least become difficult to deconvolute from diffusion artefacts resulting from the experimental method. Direct, unmediated electrochemical control is therefore an important tool for the study of redox proteins and enzymes. The technique of protein film electrochemistry (PFE) employs electrode-immobilized redox proteins, in such a way that electrons are transferred directly to or from the redox cofactors within the protein as the electrode is polarized at a series of potentials.17,18,19 PFE is of particular value for the study of oxidation or reduction reactions catalyzed by redox enzymes, as interfacial electron transfer can be achieved at a very high rate. For example, the electrocatalytic turnover rate of the nickel-iron ([NiFe]) hydrogenase from Allochromatium vinosum was measured by PFE as ca 1,000 - 10,000 s&#8722;1 for H2 oxidation.20&#160;The electrode potential acts as a trigger to turn catalysis &#39;on&#39; or &#39;off&#39;, and the electrocatalytic current reports on enzyme activity. PFE is therefore a valuable method for analyzing the reactivity of complex enzymes that depend intimately on potential, such as reactions of the di-iron active site of [FeFe]-hydrogenases with CO and O2,21 or potential-induced inactivation reactions of hydrogenases,22 carbon monoxide dehydrogenase,23 and other complex redox enzymes.24
The principal barrier to combining spectroscopic techniques with the direct electrochemical control afforded by PFE arises from the low surface coverage of redox enzymes, on the order of 1-2 pmol cm-2 for the [NiFe] hydrogenase from A. vinosum,20 relative to in situ surface science studies of small molecule adsorbates on bulk metal electrodes. This presents a challenge for the sensitivity of the spectroscopic measurement. Several spectroelectrochemical methods have been reported for studying immobilized redox proteins at a range of different electrodes: UV-visible spectroscopy at transparent metal oxide electrodes;25,26,27 fluorescence spectroscopy at gold electrodes;28,29 surface enhanced infrared absorption (SEIRA) spectroscopy at gold electrodes;30,31,32,33 and surface enhanced Raman spectroscopic techniques, principally at silver electrodes.34,35
Here we describe a method for coupling PFE with IR spectroscopy, in a technique known as protein film infrared electrochemistry (PFIRE).36 The PFIRE method studies redox enzymes immobilized on a high surface area carbon working electrode in conjunction with an attenuated total reflectance IR (ATR-IR) geometry, exploiting the ease of adsorption of a range of proteins onto carbon surfaces. IR spectroscopy is useful in studies of redox enzymes and proteins as many small molecules, ligands and cofactors have diagnostic absorbances that can be used to assess reactivity, binding, inhibition and redox state. Examples include binding of NO to iron-sulfur centers,37 study of flavoproteins,38,39,40 small molecule binding to heme centers etc.41 The ATR-IR geometry allows construction of an optimized three-electrode (spectro)electrochemical cell42 and therefore provides excellent electrochemical control. Solution resistance and potential drift are minimized by placing a reference electrode close to the working electrode. High surface area counter electrodes are used that are compatible with high electrocatalytic currents produced by fast enzyme turnover at the working electrode. Flow of solution through the spectroelectrochemical cell allows facile control over the concentration of substrates, inhibitors and pH.36,43,44 The PFIRE method therefore allows IR spectra to be recorded in situ during sustained enzyme electrocatalysis.36,44 PFIRE is also capable of providing chemical information in the absence of catalytic current,43 in contrast to PFE where it can be difficult to extract information from non-catalytic processes in redox enzymes.45,46
We have demonstrated the PFIRE method for study of electrocatalytic H2 oxidation by [NiFe] hydrogenases, which contain intrinsic CO and CN ligands coordinated to Fe at a bimetallic active site.36,43,44 [NiFe] hydrogenases are therefore particularly well suited to study by PFIRE. The PFIRE method provides information about the species that are present during steady-state turnover, and therefore provides crucial mechanistic insight in addition to the wealth of literature on IR spectroscopy of hydrogenases without experimental control over turnover.47,48 Dyer and co-workers have employed time-resolved IR methods to study of NiFe hydrogenases,49,50,51 using a light trigger to either apply a small negative potential step (via use of solution mediators and a &#39;caged&#39; electron source) or photolyse a bound hydride. Although the PFIRE method cannot, at present, provide time resolution to match these measurements,40 it does allow study of both reductive and oxidative catalytic processes, accessed at a range of well-defined potentials and free from mass transport limitations.
The PFIRE method is distinct from SEIRA studies of redox proteins, which also use an ATR-IR geometry and employ a nanoscale-roughened metal electrode to enhance the IR absorbance of molecules adsorbed on the electrode surface.30 SEIRA is an extremely valuable technique for studying membrane proteins, in particular adsorbed on or within mimetic membrane architectures,32 but the need for a metal electrode can limit the substrate and inhibitor scope due to reactivity of the electrode support towards small molecules such as CO, CN-, CO2 etc. Proton reduction and self-assembled monolayer desorption can be problematic on metal surfaces at very negative potentials,1,52 although enzyme electrocatalysis on unprotected metal electrodes has been reported.53,54 A disadvantage of PFIRE relative to SEIRA is the relative difficulty of incorporating membrane proteins in native or mimetic membrane architectures. However, the relative chemical inertness of the carbon electrodes to competing small molecule activation reactions makes PFIRE an excellent technique for studying enzyme electrocatalysis, particularly in the low-potential domain relevant to biological redox processes such as proton reduction by hydrogenases.1,43
The aim of this article is to introduce the PFIRE method as a technique for studying electrode-immobilized redox proteins, using NiFe hydrogenase 1 (Hyd1) from Escherichia coli as an example. Considerations of sample preparation, the requirement for good substrate flow, and data handling are discussed. PFIRE is a broadly applicable technique, well-suited to studying any redox protein (with characteristic IR absorbances) that can be adsorbed onto carbon electrodes, either directly or using surface modification, such that it can exchange electrons with the electrode.

<table><tbody><tr><td>Spectrometer</td><td>Agilent</td><td>680-IR</td><td>with an external MCT detector</td></tr><tr><td>ATR accessory</td><td>Pike Technologies</td><td>GladiATR</td><td>Customised for use with a 5-reflection Si IRE</td></tr><tr><td>Glovebox</td><td>Glove Box Technology Ltd.</td><td>N/A</td><td>Custom designed &#039;wet&#039; and &#039;dry&#039; box for anaerobic sample handling and measurement</td></tr><tr><td>KBr window</td><td>Crystran</td><td>Custom</td><td>To allow coupling of the glovebox with the external beam of the FTIR spectrometer</td></tr><tr><td>Additional optics</td><td>Agilent</td><td>N/A</td><td>Components from a PM-IRRAS accessory</td></tr><tr><td>Silicon IRE</td><td>Crystal GmbH</td><td>Custom</td><td>Trapezoidal: 8.4 mm x 5 mm (large face), 1 mm thickness, ca 39 degree face angle</td></tr><tr><td>Potentiostat</td><td>Metrohm</td><td>Autolab PGSTAT 128N</td><td></td></tr><tr><td>Nova 10.1</td><td>Metrohm</td><td></td><td>Software for controlling the potentiostat</td></tr><tr><td>Peristaltic pump</td><td>Williamson Manufacturing Company Ltd</td><td>QL-1000-024-300</td><td></td></tr><tr><td>Pt wire</td><td>Surepure Chemetals</td><td>3272</td><td>99.95% Pure Platinum Wire, 0.014 inch Diameter</td></tr><tr><td>Carbon rod</td><td>WH Smith</td><td>30729209</td><td>0.7 mm HB pencil lead</td></tr><tr><td>Carbon black</td><td>Cabot Corporation</td><td>Black Pearls 2000</td><td></td></tr><tr><td>Ultrasonic bath</td><td>Ultrawave</td><td>U100</td><td>35 W</td></tr><tr><td>Centrifugal filter</td><td>Merck Millipore</td><td>UFC5050BK</td><td>Amicon Ultra, 50 KDa MW cutoff</td></tr><tr><td>Microcentrifuge</td><td>Eppendorf</td><td>5452000018</td><td>MiniSpin</td></tr><tr><td>NaCl</td><td>Sigma</td><td>71376</td><td></td></tr><tr><td>H2SO4</td><td>Fisher</td><td>S/9240/PB17</td><td></td></tr><tr><td>HNO3</td><td>Fisher</td><td>N/2300/PB17</td><td></td></tr><tr><td>silicone sealant</td><td>Cow Corning Toray Co., Ltd.</td><td>SE 4486</td><td></td></tr><tr><td>Carbon paper</td><td>Toray</td><td>TGP-H-030</td><td></td></tr><tr><td>H2 gas</td><td>BOC</td><td></td><td></td></tr><tr><td>N2 gas</td><td>BOC</td><td></td><td></td></tr><tr><td>OriginPro 2015</td><td>OriginLab</td><td></td><td>Data analysis/graphing software</td></tr><tr><td>Resolutions Pro 4.0</td><td>Varian</td><td></td><td>Software for controlling FTIR spectrometer</td></tr></tbody></table>

protein film infrared electrochemistry demonstrated for study of h2 oxidation by a [nife] hydrogenase

Understanding the chemistry of redox proteins demands methods that provide precise control over redox centers within the protein. The technique of protein film electrochemistry, in which a protein is immobilized on an electrode surface such that the electrode replaces physiological electron donors or acceptors, has provided functional insight into the redox reactions of a range of different proteins. Full chemical understanding requires electrochemical control to be combined with other techniques that can add additional structural and mechanistic insight. Here we demonstrate a technique, protein film infrared electrochemistry, which combines protein film electrochemistry with infrared spectroscopic sampling of redox proteins. The technique uses a multiple-reflection attenuated total reflectance geometry to probe a redox protein immobilized on a high surface area carbon black electrode. Incorporation of this electrode into a flow cell allows solution pH or solute concentrations to be changed during measurements. This is particularly powerful in addressing redox enzymes, where rapid catalytic turnover can be sustained and controlled at the electrode allowing spectroscopic observation of long-lived intermediate species in the catalytic mechanism. We demonstrate the technique with experiments on E. coli hydrogenase 1 under turnover (H2 oxidation) and non-turnover conditions.

Watch this Scientific Journal Video about Protein Film Infrared Electrochemistry Demonstrated for Study of H2 Oxidation by a [NiFe] Hydrogenase at JoVE.com

Protein Film Infrared Electrochemistry Demonstrated for Study of H2 Oxidation by a [NiFe] Hydrogenase

Here, we describe a technique, protein film infrared electrochemistry, which allows immobilized redox proteins to be studied spectroscopically under direct electrochemical control at a carbon electrode. Infrared spectra of a single protein sample can be recorded at a range of applied potentials and under a variety of solution conditions.

Protein Film Infrared Electrochemistry Demonstrated for Study of H2 Oxidation by a [NiFe] Hydrogenase

Understanding the chemistry of redox proteins demands methods that provide precise control over redox centers within the protein. The technique of protein ...

protein-film-infrared-electrochemistry-demonstrated-for-study-h2

Research

JoVE Journal

Biochemistry

12.2K Views.  University of Oxford, Inorganic Chemistry Laboratory. Here, we describe a technique, protein film infrared electrochemistry, which allows immobilized redox proteins to be studied spectroscopically under direct electrochemical control at a carbon electrode. Infrared spectra of a single protein sample can be recorded at a range of applied potentials and under a variety of solution conditions.

Video: Protein Film Infrared Electrochemistry Demonstrated for Study of H2 Oxidation by a [NiFe] Hydrogenase

Probing and Mapping Electrode Surfaces in Solid Oxide Fuel Cells

Solid oxide fuel cells (SOFCs) are potentially the most efficient and cost-effective solution to utilization of a wide variety of fuels beyond hydrogen 1-7. The performance of SOFCs and the rates of many chemical and energy transformation processes in energy storage and conversion devices in general are limited primarily by charge and mass transfer along electrode surfaces and across interfaces. Unfortunately, the mechanistic understanding of these processes is still lacking, due largely to the difficulty of characterizing these processes under in situ conditions. This knowledge gap is a chief obstacle to SOFC commercialization. The development of tools for probing and mapping surface chemistries relevant to electrode reactions is vital to unraveling the mechanisms of surface processes and to achieving rational design of new electrode materials for more efficient energy storage and conversion2. Among the relatively few in situ surface analysis methods, Raman spectroscopy can be performed even with high temperatures and harsh atmospheres, making it ideal for characterizing chemical processes relevant to SOFC anode performance and degradation8-12. It can also be used alongside electrochemical measurements, potentially allowing direct correlation of electrochemistry to surface chemistry in an operating cell. Proper in situ Raman mapping measurements would be useful for pin-pointing important anode reaction mechanisms because of its sensitivity to the relevant species, including anode performance degradation through carbon deposition8, 10, 13, 14 ("coking") and sulfur poisoning11, 15 and the manner in which surface modifications stave off this degradation16. The current work demonstrates significant progress towards this capability. In addition, the family of scanning probe microscopy (SPM) techniques provides a special approach to interrogate the electrode surface with nanoscale resolution. Besides the surface topography that is routinely collected by AFM and STM, other properties such as local electronic states, ion diffusion coefficient and surface potential can also be investigated17-22. In this work, electrochemical measurements, Raman spectroscopy, and SPM were used in conjunction with a novel test electrode platform that consists of a Ni mesh electrode embedded in an yttria-stabilized zirconia (YSZ) electrolyte. Cell performance testing and impedance spectroscopy under fuel containing H2S was characterized, and Raman mapping was used to further elucidate the nature of sulfur poisoning. In situ Raman monitoring was used to investigate coking behavior. Finally, atomic force microscopy (AFM) and electrostatic force microscopy (EFM) were used to further visualize carbon deposition on the nanoscale. From this research, we desire to produce a more complete picture of the SOFC anode.

We present a unique platform for characterizing electrode surfaces in solid oxide fuel cells (SOFCs) that allows simultaneous performance of multiple characterization techniques (e.g. in situ Raman spectroscopy and scanning probe microscopy alongside electrochemical measurements). Complementary information from these analyses may help to advance toward a more profound understanding of electrode reaction and degradation mechanisms, providing insights into rational design of better materials for SOFCs.

Solid oxide fuel cells (SOFCs) are potentially the most efficient and cost-effective solution to utilization of a wide variety of fuels beyond hydrogen ...

Engineering

Proton Transfer and Protein Conformation Dynamics in Photosensitive Proteins by Time-resolved Step-scan Fourier-transform Infrared Spectroscopy

Monitoring the dynamics of protonation and protein backbone conformation changes during the function of a protein is an essential step towards understanding its mechanism. Protonation and conformational changes affect the vibration pattern of amino acid side chains and of the peptide bond, respectively, both of which can be probed by infrared (IR) difference spectroscopy. For proteins whose function can be repetitively and reproducibly triggered by light, it is possible to obtain infrared difference spectra with (sub)microsecond resolution over a broad spectral range using the step-scan Fourier transform infrared technique. With ~102-103 repetitions of the photoreaction, the minimum number to complete a scan at reasonable spectral resolution and bandwidth, the noise level in the absorption difference spectra can be as low as ~10-4, sufficient to follow the kinetics of protonation changes from a single amino acid. Lower noise levels can be accomplished by more data averaging and/or mathematical processing. The amount of protein required for optimal results is between 5-100 &#181;g, depending on the sampling technique used. Regarding additional requirements, the protein needs to be first concentrated in a low ionic strength buffer and then dried to form a film. The protein film is hydrated prior to the experiment, either with little droplets of water or under controlled atmospheric humidity. The attained hydration level (g of water / g of protein) is gauged from an IR absorption spectrum. To showcase the technique, we studied the photocycle of the light-driven proton-pump bacteriorhodopsin in its native purple membrane environment, and of the light-gated ion channel channelrhodopsin-2 solubilized in detergent.

Key steps of protein function, in particular backbone conformational changes and proton transfer reactions, often take place in the microsecond to millisecond time scale. These dynamical processes can be studied by time-resolved step-scan Fourier-transform infrared spectroscopy, in particular for proteins whose function is triggered by light.

Monitoring the dynamics of protonation and protein backbone conformation changes during the function of a protein is an essential step towards ...

Hydrophobic Salt-modified Nafion for Enzyme Immobilization and Stabilization

Over the last decade, there has been a wealth of application for immobilized and stabilized enzymes including biocatalysis, biosensors, and biofuel cells.1-3 In most bioelectrochemical applications, enzymes or organelles are immobilized onto an electrode surface with the use of some type of polymer matrix. This polymer scaffold should keep the enzymes stable and allow for the facile diffusion of molecules and ions in and out of the matrix. Most polymers used for this type of immobilization are based on polyamines or polyalcohols - polymers that mimic the natural environment of the enzymes that they encapsulate and stabilize the enzyme through hydrogen or ionic bonding. Another method for stabilizing enzymes involves the use of micelles, which contain hydrophobic regions that can encapsulate and stabilize enzymes.4,5 In particular, the Minteer group has developed a micellar polymer based on commercially available Nafion.6,7 Nafion itself is a micellar polymer that allows for the channel-assisted diffusion of protons and other small cations, but the micelles and channels are extremely small and the polymer is very acidic due to sulfonic acid side chains, which is unfavorable for enzyme immobilization. However, when Nafion is mixed with an excess of hydrophobic alkyl ammonium salts such as tetrabutylammonium bromide (TBAB), the quaternary ammonium cations replace the protons and become the counter ions to the sulfonate groups on the polymer side chains (Figure 1). This results in larger micelles and channels within the polymer that allow for the diffusion of large substrates and ions that are necessary for enzymatic function such as nicotinamide adenine dinucleotide (NAD). This modified Nafion polymer has been used to immobilize many different types of enzymes as well as mitochondria for use in biosensors and biofuel cells.8-12 This paper describes a novel procedure for making this micellar polymer enzyme immobilization membrane that can stabilize enzymes. The synthesis of the micellar enzyme immobilization membrane, the procedure for immobilizing enzymes within the membrane, and the assays for studying enzymatic specific activity of the immobilized enzyme are detailed below.

This article will describe the procedure for synthesizing a hydrophobically modified Nafion enzyme immobilization membrane and how to immobilize proteins and/or enzymes within the membrane and test their specific activity.

Over the last decade, there has been a wealth of application for immobilized and stabilized enzymes including biocatalysis, biosensors, and biofuel ...

Bioengineering

Light-driven Enzymatic Decarboxylation

Oxidoreductases belong to the most-applied industrial enzymes. Nevertheless, they need external electrons whose supply is often costly and challenging. Recycling of the electron donors NADH or NADPH requires the use of additional enzymes and sacrificial substrates. Interestingly, several oxidoreductases accept hydrogen peroxide as electron donor. While being inexpensive, this reagent often reduces the stability of enzymes. A solution to this problem is the in situ generation of the cofactor. The continuous supply of the cofactor at low concentration drives the reaction without impairing enzyme stability. This paper demonstrates a method for the light-catalyzed in situ generation of hydrogen peroxide with the example of the heme-dependent fatty acid decarboxylase OleTJE. The fatty acid decarboxylase OleTJE was discovered due to its unique ability to produce long-chain 1-alkenes from fatty acids, a hitherto unknown enzymatic reaction. 1-alkenes are widely used additives for plasticizers and lubricants. OleTJE has been shown to accept electrons from hydrogen peroxide for the oxidative decarboxylation. While addition of hydrogen peroxide damages the enzyme and results in low yields, in situ generation of the cofactor circumvents this problem. The photobiocatalytic system shows clear advantages regarding enzyme activity and yield, resulting in a simple and efficient system for fatty acid decarboxylation.

We describe a protocol for the light-catalyzed generation of hydrogen peroxide &#8212; a cofactor for oxidative transformations.

Oxidoreductases belong to the most-applied industrial enzymes. Nevertheless, they need external electrons whose supply is often costly and challenging. ...

Chemistry

Simultaneous Measurement of Superoxide/Hydrogen Peroxide and NADH Production by Flavin-containing Mitochondrial Dehydrogenases

It has been reported that mitochondria can contain up to 12 enzymatic sources of reactive oxygen species (ROS). A majority of these sites include flavin-dependent respiratory complexes and dehydrogenases that produce a mixture of superoxide (O2&#9679;-) and hydrogen peroxide (H2O2). Accurate quantification of the ROS-producing potential of individual sites in isolated mitochondria can be challenging due to the presence of antioxidant defense systems&#160;and side reactions that also form O2&#9679;-/H2O2. Use&#160;of nonspecific inhibitors that can disrupt mitochondrial bioenergetics can also compromise measurements by altering&#160;ROS release from other sites of production. Here, we present an easy method for the simultaneous measurement of H2O2 release and nicotinamide adenine dinucleotide (NADH) production by purified flavin-linked dehydrogenases. For our purposes here, we have used purified pyruvate dehydrogenase complex (PDHC) and &#945;-ketoglutarate dehydrogenase complex (KGDHC) of porcine heart origin as examples. This method allows for an accurate measure of native H2O2 release rates by individual sites of production by eliminating other potential sources of ROS and antioxidant systems. In addition, this method allows for a direct comparison of the relationship between H2O2 release and enzyme activity and the screening of the effectiveness and selectivity of inhibitors for ROS production. Overall, this approach can allow for the in-depth assessment of native rates of ROS release for individual enzymes prior to conducting more sophisticated experiments with isolated mitochondria or permeabilized muscle fiber.

Mitochondria contain several flavin-dependent enzymes that can produce reactive oxygen species (ROS). Monitoring ROS release from individual sites in mitochondria is challenging due to unwanted side reactions. We present an easy, inexpensive method for direct assessment of native rates for ROS release using purified flavoenzymes and microplate fluorometry.

It has been reported that mitochondria can contain up to 12 enzymatic sources of reactive oxygen species (ROS). A majority of these sites include ...

Iridium Oxide-reduced Graphene Oxide Nanohybrid Thin Film Modified Screen-printed Electrodes as Disposable Electrochemical Paper Microfluidic pH Sensors

A facile, controllable, inexpensive and green electrochemical synthesis of IrO2-graphene nanohybrid thin films is developed to fabricate an easy-to-use integrated paper microfluidic electrochemical pH sensor for resource-limited settings. Taking advantages from both pH meters and strips, the pH sensing platform is composed of hydrophobic barrier-patterned paper micropad (&#181;PAD) using polydimethylsiloxane (PDMS), screen-printed electrode (SPE) modified with IrO2-graphene films and molded acrylonitrile butadiene styrene (ABS) plastic holder. Repetitive cathodic potential cycling was employed for graphene oxide (GO) reduction which can completely remove electrochemically unstable oxygenated groups and generate a 2D defect-free homogeneous graphene thin film with excellent stability and electronic properties. A uniform and smooth IrO2 film in nanoscale grain size is anodically electrodeposited onto the graphene film, without any observable cracks. The resulting IrO2-RGO electrode showed slightly super-Nernstian responses from pH 2-12 in Britton-Robinson (B-R) buffers with good linearity, small hysteresis, low response time and reproducibility in different buffers, as well as low sensitivities to different interfering ionic species and dissolved oxygen. A simple portable digital pH meter is fabricated, whose signal is measured with a multimeter, using high input-impedance operational amplifier and consumer batteries. The pH values measured with the portable electrochemical paper-microfluidic pH sensors were consistent with those measured using a commercial laboratory pH meter with a glass electrode.

The study demonstrates the growth of iridium oxide-reduced graphene oxide (IrO2-RGO) nanohybrid thin films on irregular and rough screen-printed carbon substrate through a green electrochemical synthesis, and their implementation as a pH sensor with a patterned paper-fluidic platform.

A facile, controllable, inexpensive and green electrochemical synthesis of IrO2-graphene nanohybrid thin films is developed to fabricate an easy-to-use ...

Synthesis and Performance Characterizations of Transition Metal Single Atom Catalyst for Electrochemical CO2 Reduction

This protocol presents both the synthesis method of the Ni single atom catalyst, and the electrochemical testing of its catalytic activity and selectivity in aqueous CO2 reduction. Different from traditional metal nanocrystals, the synthesis of metal single atoms involves a matrix material that can confine those single atoms and prevent them from aggregation. We report an electrospinning and thermal annealing method to prepare Ni single atoms dispersed and coordinated in a graphene shell, as active centers for CO2 reduction to CO. During the synthesis, N dopants play a critical role in generating graphene vacancies to trap Ni atoms. Aberration-corrected scanning transmission electron microscopy and three-dimensional atom probe tomography were employed to identify the single Ni atomic sites in graphene vacancies. Detailed setup of electrochemical CO2 reduction apparatus coupled with an on-line gas chromatography is also demonstrated. Compared to metallic Ni, Ni single atom catalyst exhibit dramatically improved CO2 reduction and suppressed H2 evolution side reaction.

Synthesis and Performance Characterizations of Transition Metal Single Atom Catalyst for Electrochemical CO2 Reduction

Here, we present a protocol for the synthesis and electrochemical testing of transition metal single atoms coordinated in graphene vacancies as active centers for selective carbon dioxide reduction to carbon monoxide in aqueous solutions.

This protocol presents both the synthesis method of the Ni single atom catalyst, and the electrochemical testing of its catalytic activity and selectivity ...

Developing Photosensitizer-Cobaloxime Hybrids for Solar-Driven H2 Production in Aqueous Aerobic Conditions

Developing photocatalytic H2 production devices is the one of the key steps for constructing a global H2-based renewable energy infrastructure. A number of photoactive assemblies have emerged where a photosensitizer and cobaloxime-based H2 production catalysts work in tandem to convert light energy into the H-H chemical bonds. However, the long-term instability of these assemblies and the need for hazardous proton sources have limited their usage. Here, in this work, we have integrated a stilbene-based organic dye into the periphery of a cobaloxime core via a distinct axial pyridine linkage. This strategy allowed us to develop a photosensitizer-catalyst hybrid structure with the same molecular framework. In this article, we have explained the detailed procedure of the synthesis of this hybrid molecule in addition to its comprehensive chemical characterization. The structural and optical studies have exhibited an intense electronic interaction between the cobaloxime core and the organic photosensitizer. The cobaloxime was active for H2 production even in the presence of water as the proton source. Here, we have developed a simple airtight system connected with an online H2 detector for the investigation of the photocatalytic activity by this hybrid complex. This photosensitizer-catalyst dyad present in the experimental setup continuously produced H2 once it was exposed in the natural sunlight. This photocatalytic H2 production by the hybrid complex was observed in aqueous/organic mixture media in the presence of a sacrificial electron donor under complete aerobic conditions. Thus, this photocatalysis measurement system along with the photosensitizer-catalyst dyad provide valuable insight for the development of next generation photocatalytic H2 production devices.

Developing Photosensitizer-Cobaloxime Hybrids for Solar-Driven H2 Production in Aqueous Aerobic Conditions

We have directly incorporated a stilbene-based organic dye into a cobaloxime core to generate a photosensitizer-catalyst dyad for photocatalytic H2 production. We have also developed a simple experimental setup for evaluating the light-driven H2 production by photocatalytic assemblies.

Developing photocatalytic H2 production devices is the one of the key steps for constructing a global H2-based renewable energy infrastructure. A number ...

Catalytic Reactions at Amine-Stabilized and Ligand-Free Platinum Nanoparticles Supported on Titania During Hydrogenation of Alkenes and Aldehydes

Ligands like amines are used in the colloidal synthesis approach to protect platinum nanoparticles (Pt NP&#39;s) from agglomeration. Normally, ligands like amines are removed by diverse pre-treatment procedures before use in heterogeneous catalysis as amines are considered as a catalyst poison. However, a possible beneficial influence of these surface modifiers on hydrogenation reactions, which is known from spectator species on metal surfaces, is often neglected.
Therefore, amine-stabilized Pt nanoparticles supported by titania (P25) were used without any pre-treatment in order to elucidate a possible influence of the ligand in liquid phase hydrogenation reactions. The catalytic activity of amine-stabilized Pt nanoparticles of two different sizes was investigated in a double-walled stirring tank reactor at 69 &#176;C to 130 &#176;C and 1 atm hydrogen pressure. The conversion of cyclohexene to cyclohexane was determined by gas chromatography (GC) and was compared to ligand-free Pt particles. All catalysts were checked before and after reaction by transmission electron spectroscopy (TEM) and X-ray photoelectron spectroscopy (XPS) for possible changes in size, shape, and ligand shell. The hydrogenation of cyclohexene in liquid phase revealed a higher conversion for amine-stabilized Pt nanoparticles on titania than the ligand-free particles. The hydrogenation of 5-methylfurfural (5-MF) was chosen for a further test reaction, since the hydrogenation of &#945;, &#946;-unsaturated aldehydes is more complex and exhibits various reaction paths. However, XPS and infrared spectroscopy (IR) proved that 5-MF acts as catalyst poison at the given reaction conditions.

This protocol shows a convenient method for comparing the catalytic properties of supported platinum catalysts, synthesized by deposition of nanosized colloids or by impregnation. The hydrogenation of cyclohexene serves as a model reaction to determine the catalytic activity of the catalysts.

Ligands like amines are used in the colloidal synthesis approach to protect platinum nanoparticles (Pt NP&#39;s) from agglomeration. Normally, ligands ...

Hydrogen Production and Utilization in a Membrane Reactor

Industrial hydrogenation consumes ~11 Mt of fossil-derived H2 gas yearly. Our group invented a membrane reactor to bypass the need to use H2 gas for hydrogenation chemistry. The membrane reactor sources hydrogen from water and drives reactions using renewable electricity. In this reactor, a thin piece of Pd separates an electrochemical hydrogen production compartment from a chemical hydrogenation compartment. The Pd in the membrane reactor acts as (i) a hydrogen-selective membrane, (ii) a cathode, and (iii) a catalyst for hydrogenation. Herein, we report the use of atmospheric mass spectrometry (atm-MS) and gas chromatography mass spectrometry (GC-MS) to demonstrate that an applied electrochemical bias across a Pd membrane enables efficient hydrogenation without direct H2 input in a membrane reactor. With atm-MS, we measured a hydrogen permeation of 73%, which enabled the hydrogenation of propiophenone to propylbenzene with 100% selectivity, as measured by GC-MS. In contrast to conventional electrochemical hydrogenation, which is limited to low concentrations of starting material dissolved in a protic electrolyte, the physical separation of hydrogen production from utilization in the membrane reactor enables hydrogenation in any solvent or at any concentration. The use of high concentrations and a wide range of solvents is particularly important for reactor scalability and future commercialization.

Membrane reactors enable hydrogenation in ambient conditions without direct H2 input. We can track the hydrogen production and utilization in these systems using atmospheric mass spectrometry (atm-MS) and gas chromatography mass spectrometry (GC-MS).

Industrial hydrogenation consumes ~11 Mt of fossil-derived H2 gas yearly. Our group invented a membrane reactor to bypass the need to use H2 gas for ...

Protein Film Infrared Electrochemistry Demonstrated for Study of H₂ Oxidation by a [NiFe] Hydrogenase

Podsumowanie

Przeglądaj więcej filmów