Covalent Attachment of FeFe Hydrogenases to Carbon Electrodes for

Aug 14, 2012 - In both cases, a surface patch of lysine residues makes it possible to favor an orientation that is efficient for fast, direct electron...
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Covalent Attachment of FeFe Hydrogenases to Carbon Electrodes for Direct Electron Transfer Carole Baffert,*,† Kateryna Sybirna,‡ Pierre Ezanno,† Thomas Lautier,§ Viviane Hajj,† Isabelle Meynial-Salles,§ Philippe Soucaille,§ Hervé Bottin,‡ and Christophe Léger† †

CNRS, Aix Marseille Université, BIP UMR 7281, IMM FR 3479, 13402 Marseille Cedex 20, France iBiTec-S SB2SM, LMB (UMR CNRS 8221), DSV, CEA, 91191 Gif-sur-Yvette, France § Université de Toulouse, INSA, UPS, INP, LISBP, INRA:UMR792, CNRS:UMR 5504, 135 Avenue de Rangueil, 31077 Toulouse, France ‡

S Supporting Information *

ABSTRACT: Direct electron transfer between enzymes and electrodes is now commonly achieved, but obtaining protein films that are very stable may be challenging. This is particularly crucial in the case of hydrogenases, the enzymes that catalyze the biological conversion between dihydrogen and protons, because the instability of the hydrogenase films may prevent the use of these enzymes as electrocatalysts of H2 oxidation and production in biofuel cells and photoelectrochemical cells. Here we show that two different FeFe hydrogenases (from Chamydomonas reinhardtii and Clostridium acetobutylicum) can be covalently attached to functionalized pyrolytic graphite electrodes using peptidic coupling. In both cases, a surface patch of lysine residues makes it possible to favor an orientation that is efficient for fast, direct electron transfer. High hydrogen-oxidation current densities are maintained for up to one week, the only limitation being the intrinsic stability of the enzyme. We also show that covalent attachment has no effect on the catalytic properties of the enzyme, which means that this strategy can also used be for electrochemical studies of the catalytic mechanism.

H

exposed at the protein surface. This provides a Y-shaped redox chain that wires the active site to the site of interaction with the redox partner at the protein surface (Figure 1C). The enzyme from D. desulf uricans (purple in Figure 1C and D) is of intermediate complexity: it houses the H-cluster and two 4Fe4S clusters, one of which is surface exposed. Examining these 3D structures, one concludes that direct electron transfer to one of the bacterial enzymes requires that at least one surface exposed cluster be close to the electrode surface, whereas in the case of the algal enzyme, interfacial electron transfer should occur between the electrode and the 4Fe4S subpart of the H-cluster, which is near the protein surface. Hydrogenases can be used for hydrogen photoproduction or for H2 oxidation in biofuel cells, where they can replace the expensive catalysts made from rare metals. The photoproduction of H2 is a natural pathway in photosynthetic organism (green algea, cyanobacteria)7 but usually with low yield and taking place only in reducing conditions. Hydrogen can also be photoproduced using hydrogenases that have been coupled to photosensitizers such as CdTe nanocrystals,8 ruthenium complexes9,10 or anodized tubular TiO2;11,12 in that case visible light is the energy source for hydrogen

ydrogenases are large and complex metalloenzymes. They are present in most microorganisms, and they catalyze a reaction that is essential and has promising technological applications: the reversible conversion between molecular hydrogen and protons. There are two main types of hydrogenases depending on metal content at the active site: NiFe hydrogenases (Figure 1B) and FeFe hydrogenases (Figure 1A). The enzymes that are part of the same family have the same active site, but the proteic structure and the number of accessory electron transferring metal clusters vary depending on which organism they come from. In the case of FeFe hydrogenases, hydrogen activation occurs at the so-called H-cluster, which consists of a [Fe2(CO)3(CN)2(dtma)] subsite (dtma=dithiomethyl amine) covalently bound to a [4Fe4S] cluster (Figure 1A). FeFe hydrogenases from three organisms have been crystallized: the bacteria Clostridium pasteurianum1 and Desulfovibrio desulf uricans2 and the green algae Chlamydomonas reinhardtii.3 We study the enzymes from C. reinhardtii and Clostridium acetobutylicum which is highly homologuous to the enzyme from C. pasteurianum. The algal enzyme is the smallest hydrogenase purified so far. Its backbone is shown in blue in Figure 1D. It has a molecular weight of 49 kDa and no cofactor other than the H-cluster, which is exposed at the surface of the protein. In contrast, the enzyme from C. pasterianum has a 200-amino acid N-terminal extension, which covalently binds 4 additional FeS clusters (total Mw 64 kDa, green in Figure 1C and D), two of which are © 2012 American Chemical Society

Received: June 29, 2012 Accepted: August 14, 2012 Published: August 14, 2012 7999

dx.doi.org/10.1021/ac301812s | Anal. Chem. 2012, 84, 7999−8005

Analytical Chemistry

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groups that are present on the surface of FeFe hydrogenase, near the surface-exposed redox center that is the ideal site for electron exchange with the electrode.

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MATERIALS AND METHODS Samples of C. acetobutylicum and C. reinhardtii FeFe hydrogenases were prepared as described in refs 33, 45, and 46. 4-carboxybenzenediazonium was prepared as described in ref 42: 4-aminobenzoicacid (2.74 g, 20 mmol) was dissolved in fluoroboric acid (48%, 14.6 g, 80 mmol) and water (20 mL). After complete dissolution, a solution of sodium nitrite (1.46 g, 21.2 mmol) in water (4 mL) was added dropwise while the reaction mixture was stirred. The solution was cooled in an ice bath, and the white precipitate was filtered and washed with cold ether to give the 4-carboxybenzenediazonium tetrafluoroborate. Further purification was carried out by reprecipitation in acetonitrile/diethylether. Nitrobenzenediazonium tetrafluoroborate was purchased from ABCR. Acetonitrile, N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide (EDC), and N-hydroxysuccinimide (NHS) were from Sigma-Aldrich. The results of four electrode modification procedures are compared in this paper. Procedure A consisted in simply painting a small amount (1 μL) of enzyme solution onto a freshly polished47 homemade rotating disk pyrolitic graphite edge electrode (RDPGE) (electrode surface ∼3 mm2). For procedures B and C, the RDPGE was functionalized with carboxylic moieties by being dipped 5 min into a solution of 4carboxybenzenediazonium (2 × 10−4 M in acetonitrile), then washed with water and ethanol. In a subsequent step, for procedure C only, 1 μL of water solution of EDC (36 mM) and 1 μL of water solution of NHS (18 mM) were painted on the electrode and let dry for 10 min; the electrode was then washed with water. Then, for both procedures (B and C), 1 μL of diluted solution of hydrogenase (from 0.02 to 4 × 10−4 mg/ mL) was dropped onto the electrode surface and let react for 5 min (Scheme SI1 in the Supporting Information).48 Using procedure D, the RDPGE was functionalized with amine groups; the procedure was analogous to that described in ref 25 except that the nitrobenzenediazonium molecules were spontaneously reduced onto the graphite electrode.49 The pyrolitic graphite edge electrode was dipped for 5 min into a solution of nitrobenzenediazonium (10−4 M in acetonitrile). Then, the electrode was transferred into a standard three electrodes electrochemical cell containing 0.1 M H2SO4 solution, and the nitro group was electrochemically reduced into an amine by cycling the electrode potential from 0.89 V to −0.76 V vs SHE at 200 mV/s. In a subsequent step, 1 μL of water solution of EDC (36 mM), 1 μL of water solution of NHS (18 mM), and 1 μL of diluted solution of hydrogenase (from 0.02 to 4 × 10−4 mg/mL) were dropped onto the electrode surface and let to react for 10 min. Before electrode modification, the enzyme samples were diluted into buffered solutions at pH 6 or 8 to reach final concentrations of about 0.5 μM (indeed, we observed that the obtained current densities were not greater using more concentrated enzyme solutions). The pKa of benzoic acid is 4.2 and that of the amine group of lysine is 10.5. The carboxyl group should therefore be deprotonated and the amine group protonated both at pH 6 and at pH 8, and indeed, the pH value seemed to make no difference to the signals we observed (this contrasts with an earlier report on the covalent attachment of a NiFe hydrogenase25). It should be noted that EDC cross-

Figure 1. Panels A and B are models of the active sites of (A) FeFe hydrogenase and (B) NiFe hydrogenase (color code: gray = C, red = O, blue = N, orange = Fe, green = Ni, yellow = S). The asterisk in panel A indicates the Fe ion where H2 binds during the catalytic cycle.4 (C) The “naked” inorganic cofactors of C. pasteurianum, including the H-cluster (rightmost), viewed in the same orientation as in panel D; the lines contour the cofactors that are present in each enzyme, using the same color code as in panel D. (D) Superposition of the backbones of the FeFe hydrogenases from C. pasteurianum (green, PDB accession code 3C8Y5), Desulfovibrio desulf uricans (purple, PDB 1HFE2) and Chlamydomonas reinhardtii (blue, PDB 3LX46).

production, and, in the most favorable case, for water splitting into H2 and O2.12 Biofuel cells could be used as low power devices (up to a few mW/m2) running from various substrates such as glucose, dioxygen, methanol and hydrogen.13 Photoelectrochemical biofuel cells where the anode was modified by hydrogenases have also been developed14−17 but some problems must be overcome such as the sensitivity of hydrogenases to the inhibitors O2 and CO. All electrochemical applications of hydrogenases require that the enzyme interacts strongly with, and quickly transfers electrons to an electrode, and many research groups have reported that certain electrode materials gently interact with these fragile proteins. NiFe hydrogenases have been immobilized on conducting surfaces using various methods: physical adsorption onto pyrolitic graphite,18−20 adsorption on gold surfaces modified by a self-assembled monolayer of thiols (SAM) or polymixin,21−24 and covalent attachment to carbon electrodes or nanotubes25−28 or to the amino group of SAM modified gold electrodes.29,30 FeFe hydrogenases have been adsorbed on pyrolitic graphite,31−33 TiO2,34 carbon nanotubes,35,36 and gold37,38 but in not a single case has the covalent attachment of a FeFe hydrogenase been reported. Here, we describe the successful covalent attachment of two distinct FeFe hydrogenases onto pyrolytic graphite modified with a diazonium that bears a carboxylate function. A carbon electrode is advantageous compared to gold, because it has a low cost and because carbon−carbon bonds are more stable than the gold− sulfur bonds formed in SAM electrodes.39 The covalent attachment of proteins onto carbon electrodes using aromatic diazoniums salts and peptidic coupling has been described for NiFe hydrogenases,25−27 glucose oxidase,40,41 horseradish peroxidase,42 laccase43 and cellobiose dehydrogenase.44 We adapted this approach to make use of the patches of amino 8000

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linking is most efficient under acidic conditions (pH 4.5)50 while the reaction of NHS ester-activated compounds with primary amines in slightly favored under alkaline conditions (pH 7.2−8.5).51 We could not test extreme pH values (below pH 4.5 and above pH 10) because the enzymes we study are unstable under these conditions. Adapting the pH value at each step could be a way to optimize carboxylate activation in the future. Protein film electrochemistry experiments (cyclic voltammetry and chronoamperometry) were carried out in a glovebox filled with N2, using the electrochemical setup and equipment previously described.52 The two-compartment electrochemical cell was kept at the desired temperature value using a water circulation system. The rotating disk pyrolytic graphite edge working electrode (RDPGE) (area A ≈ 3 mm2) was used in conjunction with an electrode rotator, a platinum wire was used as a counter electrode, and a saturated calomel electrode (SCE), located in a side arm containing 0.1 M NaCl and maintained at room temperature, was used as a reference. All potentials are quoted versus the standard hydrogen electrode (SHE), (ESHE = ESCE + 240 mV). The electrochemical cell contained a buffer mixture of MES, CHES, TAPS, HEPES, and sodium acetate (5 mM each), 1 mM EDTA, and 0.1 M NaCl; the temperature (T) and pH are indicated in each caption. We analyzed and fit the data using an in-house program called SOAS,53 which is available free and free of charge on our Web site at http://bip.cnrs-mrs.fr/bip06/software.html. Note that SOAS is being replaced by an entirely new, powerful, open program called QSOAS, which will become available soon. Check our webpages for updates.

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RESULTS Figure 2 shows the surface of the two proteins we studied. The published structure of the FeFe hydrogenase from Chlamydomonas reinhardtii6 has been obtained from a protein that lacks the 2Fe subpart of the H-cluster (rightmost in Figure 1A). Figure 2A shows a structural model where we added the 2Fe cluster to the apoprotein crystal structure. The contact potential calculated using the software pymol54 is shown in Figure 2A,B (the latter is obtained by a 180° rotation). The charge distribution is homogeneous, except for a patch of alkaline residues (pointed to by a black arrow) that is near the 4Fe4S cluster. The presence of this positive patch near the 4Fe4S cluster suggests that this region should interact preferentially with a negatively charged surface. The structure of Clostridium acetobutylicum FeFe hydrogenase has not yet been solved, and we used the SWISSMODEL software (http://swissmodel.expasy.org)55 to combine the known amino acid sequence of the enzyme from C. acetobutylicum56 and the structure of the highly homologous enzyme from Clostridium pasteurianum (PDB 3C8Y5) into the structural model shown in Figure 2C−F. From the examination of panels C and E in Figure 2, we conclude that surface residues near the 2Fe2S cluster and the distal 4Fe4S cluster of C. acetobutylicum FeFe hydrogenase are basic. The 2Fe-2S cluster and the distal 4Fe-4S cluster are on opposite sides of the protein, each embedded in their own lysine patch. Since in both enzymes, a patch of basic (mostly lysine) residues near the most exposed FeS clusters should favor the interaction with a negatively charged surface, we decided to use these lysine (or arginine) side-chains for covalent attachment to graphite electrodes modified with a carboxylate function.

Figure 2. (A) Contact potential of the modeled structure of the FeFe hydrogenase from Chlamydomonas reinhardtii (the 2Fe subcluster was added to the apoprotein crystal structure, the H-cluster can be seen through the transparent protein surface);3 (B) after 180° rotation of panel A; (C) contact potential of the modeled structure of Clostridium acetobutylicum FeFe hydrogenase; (E) after 180° rotation of panel C; (D) the “naked” inorganic cofactors of C. acetobutylicum; (F) after 180° rotation of panel E. The black arrows point to basic patches on the protein surface, those that are near the surface exposed clusters, and should interact favorably with carboxylate groups grafted onto electrode surfaces.

Figure 3 compares the catalytic cyclic voltammograms for C. acetobutylicum and C. reinhardtii hydrogenases (left and right columns, respectively), obtained after the electrode had been modified using one out of four different procedures: (A) the enzyme solution was simply painted onto a rotating disk pyrolitic graphite edge electrode (RDPGE); (B) the enzyme solution was painted onto a RDPGE modified by a benzoic acid group;49,57 (C) the enzyme solution was painted onto the RDPGE electrode modified by benzoic acid group in the presence of carbodiimide; (D) the enzyme solution was painted onto a RDPGE modified by aniline (from the spontaneous reduction of nitrobenzenediazonium at the electrode and electroreduction of the nitro group in an acidic solution) in the presence of carbodiimide. The rotating disk electrodes were spun at a high rate, to avoid any limitation by the transport of H2, so that steady-states conditions were achieved. The current magnitudes and catalytic waveshapes are similar for conditions A, B, and C, whereas condition D leads to a much lower current (for both enzymes). We also note that the shapes of the cyclic voltammograms are significantly different in 8001

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Figure 4. Normalized chronoamperograms for protein films of C. acetobutylicum (panels A and B) and C. reinhardtii (panels C and D) FeFe hydrogenases, under conditions where the enzymes oxidize H2 (E = −160 mV vs SHE, 1 atm of H2, pH = 7, T = 30 °C, ω = 3000 rpm). The enzymes were attached following procedure A (green lines, physical adsorption onto graphite), procedure B (blue lines, the electrode was modified with 4-carboxybenzenediazonium but no coupling agents were used), or procedure C (red lines, the enzyme was covalently attached to the electrode by reaction with benzoic acid at the electrode surface). The lines with identical colors illustrate the results of independent runs.

Figure 3. Cyclic voltammograms for protein films of C. acetobutylicum (left panel) and C. reinhardtii (right panel) FeFe hydrogenases (1 atm of H2, pH = 7, T = 30 °C, ω = 3000 rpm). The enzymes were attached using procedure A (panels A1 and A2, green lines, physical adsorption onto graphite), procedure B (panels B1 and B2, blue lines, the electrode was modified with 4-carboxybenzenediazonium, but no coupling agents were used), procedure C (panels C1 and C2, red lines, the enzyme was covalently attached to the electrode by reaction with benzoic acid at the electrode surface), or procedure D (panels D1 and D2, black lines, the enzyme was covalently attached to the electrode by reaction with aniline at the electrode surface). The dashed line in panel D2 shows a blank recorded with no enzyme.

exponential phases observed in at least three independent runs) are shown in Table 1. In all cases, we observed a fast decay with a time constant of about 25 min. The slower phase is characterized by a time constant of about 5−10 h if procedures A or B are used to attach the enzyme and 1 to 2 days with procedure C. The filled squares in Figure 4B,D show the current densities for C. acetobutylicum and C. reinhardtii hydrogenase modified electrodes (conditions A and C) measured at regular intervals for a few days. Between measurements, the electrode was disconnected and stored at 4 °C in pH 7 buffer in the glovebox used for the electrochemical experiments ([O2] < 1 ppm). The fact that these data points follow the trend observed on shorter time scales (fitting an exponential decay through these data points returns time constants that are consistent with the slow decay observed on shorter time scales, in panels A and C); this suggests that the decrease in current is almost the same under operation at 30 °C and upon storage at 4 °C. We compared data obtained with electrodes modified following procedure D (attachment to aniline group, black lines in Figure SI2 in the Supporting Information) and procedure C (attachment to benzoic acid group, red lines in Figure SI2 in the Supporting Information). We observed that much smaller current densities were obtained with procedure D than with procedure C. However, the current decays were also biphasic, and the best parameters obtained by fitting to eq 1 were similar to those obtained for procedure C. Since our group is mostly interested in using direct electrochemistry of enzymes for mechanistic studies,19 it was important to check that the enzyme properties are not changed by the covalent attachment (condition A versus condition C).

condition D, which we interpret as indicating that in this case, interfacial electron transfer between the enzyme and the electrode is particularly sluggish. We note that in all cases, the fact that interfacial electron transfer is not very fast compared to catalytic turnover significantly broadens the wave shapes, as observed before with other hydrogenases.58 We are currently examining which models can be used to describe these wave shapes and gain information about the properties of the catalytic intermediates. We tested the long-term stability of the protein films using chronoamperometry experiments, where we measure the H2 oxidation catalytic current at a constant potential (E = −160 mV vs SHE, pH 7, 30 °C) as a function of time. Figure 4 shows the results when we tested procedures A, B, and C (green, red, and blue traces, respectively). In no case did the change in current against time follow a monoexponential decay. We analyzed the current variations by fitting the data to a sum of two exponentials (noting that more complex combinations of exponential functions lead to parameter indetermination). i(t )/i(0) = a1exp(−t /τ1) + (1 − a1) exp(−t /τ2)

(1)

Since this function tends to zero, both time constants can be unambiguously determined provided the experiment lasts much longer than the shorter time constant. The results of the fits (the amplitudes a1 and a2 = 1 − a 1 and time constants of the two 8002

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Table 1. Best Parameters Obtained by Fitting the Current Decays Shown in Figure 4 to Equation 1 τ1 (min)

enzyme

procedure

a1

C. acetobutylicum

A B C A B C

0.47−0.85 0.45 0.15−0.30 0.20−0.37 0.60 0.12−0.15

C. reinhardtii

20 17 28 25 40 27

±5 ±5 ±8 ±5

a2

τ2 (h)

0.15−0.53 0.55 0.70−0.85 0.63−0.80 0.40 0.85−0.88

4.4 ± 1 7.8 50 ± 5 6.7 ± 2.5 11 22 ± 5

Table 2. Enzymes Properties for Protein Films of C. acetobutylicum and C. reinhardtii FeFe Hydrogenasesa enzyme C. acetobutylicum C. acetobutylicum C. acetobutylicum C. reinhardtii C. reinhardtii C. reinhardtii

procedure A C e g A C f g

Km (H2) (bar H2) 1.2 ± 1.1 ± 1.1b 0.59 0.5 ± 0.5 ±

kinCO (s−1 mM CO−1)

b

0.2 0.2b

0.2b 0.2b

0.24

18 ± 16 ± 20c 1.1 50 ± 65 ± 80c 19

c

2 2c

10c 10c

koutCO (s−1) −2c

1.5 × 10 1.6 × 10−2c 1.5 × 10−2c 1.8 × 10−3 2.6 × 10−2c 2.8 × 10−2c 3 × 10−2c 2.3 × 10−3

kapp(O2) (s−1 mM O2−1) −2

± 0.01 × 10 ± 0.1 × 10−2 ± 0.1 × 10−2 ± 0.1 × 10−2 ± 0.2 × 10−2

4 × 10−2d ± 2 × 10−2 5 × 10−2d ± 2 × 10−2 5.6 × 10−2d 5.1 × 10−3 6.5 ± 2d 5 ± 2d 0.22

a

The enzymes were attached using procedure A (physical adsorption onto graphite) or procedure C (the enzyme was covalently attached to the electrode by reaction with benzoic acid at the electrode surface). bThe Km values were measured following the method described in ref 60 at E = −160 mV vs SHE. cThe kinetics constants relative to CO binding and release were determined as described in ref 61 at E = −160 mV vs SHE. d app k (O2) is the apparent bimolecular rate constant relative to the reaction with O2 defined in ref 32 determined at E = 40 mV vs SHE for C. acetobutylicum hydrogenase and E = 90 mV vs SHE for C. reinhardtii hydrogenase. All experiments (but those carried out by Goldet et al.62) were carried out at T = 30 °C, pH = 7, ω = 3000 rpm, 1 bar H2. eData from ref 59. fData from ref 33. gData from ref 62. T = 10 °C, pH = 6 (note that the low temperature used in these experiments explains the lower values of the Michaelis and rate constants).

reveals that the adsorption of the enzyme leads to two different adsorption modes. In procedure C, the enzyme is dropped onto the electrode modified with carboxylate groups and coupling agents (EDC/NHS) are used. This leads to the most stable film, with a slower time constant of the order of 1−2 days. This enhanced stability of the film is likely to reveal that covalent bonds have indeed been formed between the electrode and the enzyme. Once the fast phase is over, the current decreases only 2 or 5% per hour (C. acetobutylicum or C. reinhardtii hydrogenase, respectively). This loss of current may be due to the slow degradation or denaturation of the enzyme during catalysis or storage. However, we have observed that electrodes modified with the enzymes from both C. acetobutylicum and C. reinhardtii maintain current for at least 1 week upon storage at 4 °C in an anaerobic glovebox. Concentrated enzyme samples stored at 4 °C in the glovebox maintained activity for weeks, and it therefore appears that the presence of the electrode somewhat destabilizes the protein. An electrode that gently interacts with and quickly transfers electrons to an enzyme should mimic its natural redox partner. In vivo, FeFe hydrogenases exchange electrons with plant type ferredoxins (which are small proteins containing a 2Fe2S cluster) or flavodoxins.63−65 In ref 65 it was proved that lysine side chains that are exposed at the surface of C. reinhardtii hydrogenase and acidic residues of ferredoxin PetF are involved in complex formation and electron transfer. We conclude that benzoic acid modified electrodes reproduce the acidic surface of ferredoxins and favor the orientation onto the electrode surface that allows fast and direct electron transfer. This is consistent with the observation that modifying the electrode with amino groups before forming a covalent bond (procedure D) resulted in smaller current densities (Figure SI2 in the Supporting Information) and slower interfacial electron transfer (according to the catalytic waveshapes shown in Figure 3).

For this purpose, we determined the Michaelis constant (Km) for hydrogen, the rates of inhibition by O2 and CO, and the rate of CO release, using methods developed and described previously.33,32,59 Table 2 summarizes the results we obtained.

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DISCUSSION To elaborate a strategy for the immobilization of FeFe hydrogenases, we analyzed the surface charges and the surface exposed residues that are near the exposed FeS clusters (Figure 2). We predicted that a negatively charged electrode surface should favor direct electron transfer to and from hydrogenase. In attempts to attach two distinct FeFe hydrogenases to carbon electrodes, we performed electrochemical experiments using four electrode modification procedures. We examined the magnitude of the catalytic current for H2 oxidation measured under identical conditions, the stability of this current (Figure 4), and the shape of the catalytic voltammograms (Figure 3), which embeds information about the rate of interfacial electron transfer between the electrode and the enzyme.19,58 The stability of the protein films was assessed by carrying out chronoamperometry experiments. The current variations (Figure 4) were fitted to biexponential decays (eq 1). Under all conditions of electrode modification by FeFe hydrogenase, a fast decay of current (time constant of about 25 min) was observed (Table 1). We consider as likely that this corresponds to the desorption of enzyme molecules that interact weakly with the graphite electrode. When we used procedures A or B, the enzyme was adsorbed onto rotating disk pyrolitic graphite edge (RDPGE) electrodes (with or without previous modification by a benzoic acid group from spontaneous reduction of 4-carboxybenzenediazonium) and no covalent bond could be formed. The slow decay (slower time constant ranging from 5 to 10 h) must be due to enzyme molecules that interact strongly with the electrode surface. We believe that the biexponential decay observed when procedure A or B is used 8003

dx.doi.org/10.1021/ac301812s | Anal. Chem. 2012, 84, 7999−8005

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(6) Mulder, D. W.; Boyd, E. S.; Sarma, R.; Lange, R. K.; Endrizzi, J. A.; Broderick, J. B.; Peters, J. W. Nature 2010, 465 (7295), 248−251. (7) Stripp, S. T.; Happe, T. Dalton Trans. 2009, No. 45, 9960−9969. (8) Brown, K. A.; Dayal, S.; Ai, X.; Rumbles, G.; King, P. W. J. Am. Chem. Soc. 2010, 132 (28), 9672−9680. (9) Reisner, E.; Powell, D. J.; Cavazza, C.; Fontecilla-Camps, J. C.; Armstrong, F. A. J. Am. Chem. Soc. 2009, 131 (51), 18457−18466. (10) Zadvornyy, O. A.; Lucon, J. E.; Gerlach, R.; Zorin, N. A.; Douglas, T.; Elgren, T. E.; Peters, J. W. J. Inorg. Biochem. 2012, 106 (1), 151−155. (11) Joo, H.; Bae, S.; Kim, C.; Kim, S.; Yoon, J. Sol. Energy Mater. Sol. Cells 2009, 93 (9), 1555−1561. (12) Bae, S.; Shim, E.; Yoon, J.; Joo, H. J. Power Sources 2008, 185 (1), 439−444. (13) Davis, F.; Higson, S. P. J. Biosens. Bioelectron. 2007, 22 (7), 1224−1235. (14) Hambourger, M.; Gervaldo, M.; Svedruzic, D.; King, P. W.; Gust, D.; Ghirardi, M.; Moore, A. L.; Moore, T. A. J. Am. Chem. Soc. 2008, 130 (6), 2015−2022. (15) Vincent, K. A.; Cracknell, J. A.; Parkin, A.; Armstrong, F. A. Dalton Trans. 2005, No. 21, 3397−3403. (16) Vincent, K. A.; Cracknell, J. A.; Lenz, O.; Zebger, I.; Friedrich, B.; Armstrong, F. A. Proc. Natl. Acad. Sci. U.S.A. 2005, 102 (47), 16951−16954. (17) Vincent, K. A.; Cracknell, J. A.; Clark, J. R.; Ludwig, M.; Lenz, O.; Friedrich, B.; Armstrong, F. A. Chem. Commun. 2006, No. 48, 5033−5035. (18) Armstrong, F. A.; Belsey, N. A.; Cracknell, J. A.; Goldet, G.; Parkin, A.; Reisner, E.; Vincent, K. A.; Wait, A. F. Chem. Soc. Rev. 2009, 38 (1), 36−51. (19) Leger, C.; Bertrand, P. Chem. Rev. 2008, 108 (7), 2379−2438. (20) Vincent, K. A.; Li, X.; Blanford, C. F.; Belsey, N. A.; Weiner, J. H.; Armstrong, F. A. Nat. Chem. Biol. 2007, 3 (12), 760−761. (21) Hoeben, F. J. M.; Heller, I.; Albracht, S. P. J.; Dekker, C.; Lemay, S. G.; Heering, H. A. Langmuir 2008, 24 (11), 5925−5931. (22) Hoeben, F. J. M.; Meijer, F. S.; Dekker, C.; Albracht, S. P. J.; Heering, H. A.; Lemay, S. G. ACS Nano 2008, 2 (12), 2497−2504. (23) Millo, D.; Pandelia, M. E.; Utesch, T.; Wisitruangsakul, N.; Mroginski, M. A.; Lubitz, W.; Hildebrandt, P.; Zebger, I. J. Phys. Chem. B 2009, 113 (46), 15344−15351. (24) Ciaccafava, A.; Infossi, P.; Ilbert, M.; Guiral, M.; Lecomte, S.; Giudici-Orticoni, M. T.; Lojou, E. Angew. Chem., Int. Ed. 2012, 51 (4), 953−956. (25) Rudiger, O.; Abad, J. M.; Hatchikian, E. C.; Fernandez, V. M.; De Lacey, A. L. J. Am. Chem. Soc. 2005, 127 (46), 16008−16009. (26) Alonso-Lomillo, M. A.; Rudiger, O.; Maroto-Valiente, A.; Velez, M.; Rodriguez-Ramos, I.; Munoz, F. J.; Fernandez, V. M.; De Lacey, A. L. Nano Lett. 2007, 7 (6), 1603−1608. (27) Lojou, E.; Luo, X.; Brugna, M.; Candoni, N.; Dementin, S.; Giudici-Orticoni, M. T. J. Biol. Inorg. Chem. 2008, 13 (7), 1157−1167. (28) Krishnan, S.; Armstrong, F. A. Chem. Sci. 2012, 3 (4), 1015− 1023. (29) Luo, X. J.; Brugna, M.; Tron-Infossi, P.; Giudici-Orticoni, M. T.; Lojou, E. J. Biol. Inorg. Chem. 2009, 14 (8), 1275−1288. (30) Rudiger, O.; Gutierrez-Sanchez, C.; Olea, D.; Pereira, I. A. C.; Velez, M.; Fernandez, V. M.; De Lacey, A. L. Electroanalysis 2010, 22 (7−8), 776−783. (31) Parkin, A.; Cavazza, C.; Fontecilla-Camps, J. C.; Armstrong, F. A. J. Am. Chem. Soc. 2006, 128 (51), 16808−16815. (32) Baffert, C.; Demuez, M.; Cournac, L.; Burlat, B.; Guigliarelli, B.; Bertrand, P.; Girbal, L.; Leger, C. Angew. Chem., Int. Ed. 2008, 47 (11), 2052−2054. (33) Baffert, C.; Bertini, L.; Lautier, T.; Greco, C.; Sybirna, K.; Ezanno, P.; Etienne, E.; Soucaille, P.; Bertrand, P.; Bottin, H.; MeynialSalles, I.; De Gioia, L.; Leger, C. J. Am. Chem. Soc. 2011, 133 (7), 2096−2099. (34) Morra, S.; Valetti, F.; Sadeghi, S. J.; King, P. W.; Meyer, T.; Gilardi, G. Chem. Commun. 2011, 47 (38), 10566−10568.

In clostridial hydrogenases, the electron transfer chain has a very unusual Y shape (Figure 2D,F) and it is unknown which surface exposed cluster is the site of interaction with the natural partner. We have observed that basic residues surround both the distal 4Fe4S and the 2Fe2S cluster in the enzyme from C. acetobutylicum. However, Figure SI3 in the Supporting Information shows that the contact potential of Clostridium pasteurianum FeFe hydrogenase is significantly different from that of C. acetobutylicum hydrogenase (Figure 2C,E). A major difference is the absence of a positive patch near the 2Fe2S cluster in the enzyme from C. pasteurianum. This suggests that, in this enzyme, either only the 4Fe4S cluster exchanges electrons with the ferredoxin or the 2Fe2S cluster is the site of interaction with another redox partner with a more neutral surface. We used various electrochemical methods to measure a number of kinetic parameters and compared the values obtained with the enzymes simply adsorbed or covalently attached to the electrode surface. No significant differences were found (Table 2), suggesting that the attachment method we propose can be used in experiments that aim at investigating the catalytic properties of the enzyme, without having to correct the decrease in current that is due to enzyme desorption.53 Overall, we conclude that the method we presented for covalent attachment of FeFe hydrogenases to carbon electrodes via 4-carboxybenzenediazonium electrode modification and peptidic coupling is efficient and can be used for mechanistic studies and, possibly, for future biotechnological applications.

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ASSOCIATED CONTENT

S Supporting Information *

Scheme of covalent attachment principle, comparison of current stability using procedures C and D, and surface contact of structure of C. pasteurianum FeFe hydrogenase. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: carole.baff[email protected]. Address: CNRS, Aix Marseille Université, BIP UMR 7281, IMM FR 3479, 31 Chemin Joseph Aiguier, 13402 Marseille Cedex 20, France. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Dr. Elisabeth Lojou for helpful discussions. Our work is funded by the CNRS, Université Aix-Marseille, ANR, region Provence Alpes Côte d’Azur, Ville de Marseille. We acknowledge support from the “Pôle de Compétitivité Capénergies”. K.S. was supported by the European Commission (Grant STRP SOLAR-H2 212508) then by ANR (Algo H2).

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REFERENCES

(1) Peters, J. W. Science 1999, 283 (5410), 2102−2102. (2) Nicolet, Y.; Piras, C.; Legrand, P.; Hatchikian, C. E.; FontecillaCamps, J. C. Struct. Fold. Des. 1999, 7 (1), 13−23. (3) Mulder, D. W.; Shepard, E. M.; Meuser, J. E.; Joshi, N.; King, P. W.; Posewitz, M. C.; Broderick, J. B.; Peters, J. W. Structure 2011, 19 (8), 1038−1052. (4) Greco, C.; Bruschi, M.; Fantucci, P.; Ryde, U.; De Gioia, L. ChemPhysChem 2011, 12 (17), 3376−3382. (5) Pandey, A. S.; Harris, T. V.; Giles, L. J.; Peters, J. W.; Szilagyi, R. K. J. Am. Chem. Soc. 2008, 130 (13), 4533−4540. 8004

dx.doi.org/10.1021/ac301812s | Anal. Chem. 2012, 84, 7999−8005

Analytical Chemistry

Article

(35) Svedruzic, D.; Blackburn, J. L.; Tenent, R. C.; Rocha, J. D. R.; Vinzant, T. B.; Heben, M. J.; King, P. W. J. Am. Chem. Soc. 2011, 133 (12), 4299−4306. (36) McDonald, T. J.; Svedruzic, D.; Kim, Y. H.; Blackburn, J. L.; Zhang, S. B.; King, P. W.; Heben, M. J. Nano Lett. 2007, 7 (11), 3528−3534. (37) Krassen, H.; Stripp, S.; von Abendroth, G.; Ataka, K.; Happe, T.; Heberle, J. J. Biotechnol. 2009, 142 (1), 3−9. (38) Krassen, H.; Stripp, S. T.; Bohm, N.; Berkessel, A.; Happe, T.; Ataka, K.; Heberle, J. Eur. J. Inorg. Chem. 2011, No. 7, 1138−1146. (39) D’Amours, M.; Belanger, D. J. Phys. Chem. B 2003, 107 (20), 4811−4817. (40) Bourdillon, C.; Delamar, M.; Demaille, C.; Hitmi, R.; Moiroux, J.; Pinson, J. J. Electroanal. Chem. 1992, 336 (1−2), 113−123. (41) Pellissier, M.; Barriere, F.; Downard, A. J.; Leech, D. Electrochem. Commun. 2008, 10 (6), 835−838. (42) Polsky, R.; Harper, J. C.; Dirk, S. M.; Arango, D. C.; Wheeler, D. R.; Brozik, S. M. Langmuir 2007, 23 (2), 364−366. (43) Sosna, M.; Chretien, J. M.; Kilburn, J. D.; Bartlett, P. N. Phys. Chem. Chem. Phys. 2010, 12 (34), 10018−10026. (44) Tasca, F.; Harreither, W.; Ludwig, R.; Gooding, J. J.; Gorton, L. Anal. Chem. 2011, 83 (8), 3042−3049. (45) Sybirna, K.; Antoine, T.; Lindberg, P.; Fourmond, V.; Rousset, M.; Mejean, V.; Bottin, H. BMC Biotechnol. 2008, 8, 73. (46) Girbal, L.; von Abendroth, G.; Winkler, M.; Benton, P. M. C.; Meynial-Salles, I.; Croux, C.; Peters, J. W.; Happe, T.; Soucaille, P. Appl. Environ. Microbiol. 2005, 71 (5), 2777−2781. (47) Blanford, C. F.; Armstrong, F. A. J. Solid State Electrochem. 2006, 10 (10), 826−832. (48) Jiang, K. Y.; Schadler, L. S.; Siegel, R. W.; Zhang, X. J.; Zhang, H. F.; Terrones, M. J. Mater. Chem. 2004, 14 (1), 37−39. (49) Lehr, J.; Williamson, B. E.; Downard, A. J. J. Phys. Chem. C 2011, 115 (14), 6629−6634. (50) Nakajima, N.; Ikada, Y. Bioconjugate Chem. 1995, 6 (1), 123− 130. (51) Mattson, G.; Conklin, E.; Desai, S.; Nielander, G.; Savage, M. D.; Morgensen, S. Mol. Biol. Rep. 1993, 17 (3), 167−183. (52) Liebgott, P. P.; Leroux, F.; Burlat, B.; Dementin, S.; Baffert, C.; Lautier, T.; Fourmond, V.; Ceccaldi, P.; Cavazza, C.; Meynial-Salles, I.; Soucaille, P.; Fontecilla-Camps, J. C.; Guigliarelli, B.; Bertrand, P.; Rousset, M.; Leger, C. Nat. Chem. Biol. 2010, 6 (1), 63−70. (53) Fourmond, V.; Lautier, T.; Baffert, C.; Leroux, F.; Liebgott, P. P.; Dementin, S.; Rousset, M.; Arnoux, P.; Pignol, D.; Meynial-Salles, I.; Soucaille, P.; Bertrand, P.; Leger, C. Anal. Chem. 2009, 81 (8), 2962−2968. (54) The PyMOL Molecular Graphics System; Schrödinger, LLC: Portland, OR, 2009. (55) Schwede, T.; Kopp, J. r.; Guex, N.; Peitsch, M. Nucleic Acids Res. 2003, 31 (13), 3381−3385. (56) Gorwa, M. F.; Croux, C.; Soucaille, P. J. Bacteriol. 1996, 178 (9), 2668−2675. (57) Cullen, R. J.; Jayasundara, D. R.; Soldi, L.; Cheng, J. J.; Dufaure, G.; Colavita, P. E. Chem. Mater. 2012, 24 (6), 1031−1040. (58) Leger, C.; Jones, A. K.; Albracht, S. P. J.; Armstrong, F. A. J. Phys. Chem. B 2002, 106 (50), 13058−13063. (59) Lautier, T.; Ezanno, P.; Baffert, C.; Fourmond, V.; Cournac, L.; Fontecilla-Camps, J. C.; Soucaille, P.; Bertrand, P.; Meynial-Salles, I.; Leger, C. Faraday Discuss. 2011, 148, 385−407. (60) Leger, C.; Dementin, S.; Bertrand, P.; Rousset, M.; Guigliarelli, B. J. Am. Chem. Soc. 2004, 126 (38), 12162−12172. (61) Leroux, F.; Dementin, S.; Burlatt, B.; Cournac, L.; Volbeda, A.; Champ, S.; Martin, L.; Guigliarelli, B.; Bertrand, P.; Fontecilla-Camps, J.; Rousset, M.; Leger, C. Proc. Natl. Acad. Sci. U.S.A. 2008, 105 (32), 11188−11193. (62) Goldet, G.; Brandmayr, C.; Stripp, S. T.; Happe, T.; Cavazza, C.; Fontecilla-Camps, J. C.; Armstrong, F. A. J. Am. Chem. Soc. 2009, 131 (41), 14979−14989. (63) Guerrini, O.; Burlat, B.; Leger, C.; Guigliarelli, B.; Soucaille, P.; Girbal, L. Curr. Microbiol. 2008, 56 (3), 261−267.

(64) Demuez, M.; Cournac, L.; Guerrini, O.; Soucaille, P.; Girbal, L. FEMS Microbiol. Lett. 2007, 275 (1), 113−121. (65) Winkler, M.; Kuhlgert, S.; Hippler, M.; Happe, T. J. Biol. Chem. 2009, 284 (52), 36620−36627.

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