Electrochemically Induced Far-Infrared Difference Spectroscopy on

Jan 29, 2013 - Electrochemically Induced Far-Infrared Difference Spectroscopy on Metalloproteins Using Advanced Synchrotron Technology...
1 downloads 4 Views 2MB Size
Download PDF
Article pubs.acs.org/ac

Electrochemically Induced Far-Infrared Difference Spectroscopy on Metalloproteins Using Advanced Synchrotron Technology Nicolas Vita,†,‡,§,∥ Jean-Blaise Brubach,*,∥ Rainer Hienerwadel,‡,§,▽ Nicolas Bremond,†,‡,§ Dorothée Berthomieu,¶ Pascale Roy,∥ and Catherine Berthomieu*,†,‡,§ †

Lab Interactions Protein Metal, Commissariat à l’Energie Atomique (CEA), DSV, IBEB, Saint-Paul-lez-Durance, F-13108, France Centre National de la Recherche Scientifique, UMR Biol Veget et Microbiol Environ, Saint-Paul-lez-Durance, F-13108, France § Aix-Marseille Université, Saint-Paul-lez-Durance, F-13108, France ∥ Société Civile Synchrotron SOLEIL, L’Orme des Merisiers, St-Aubin BP48, 91192 Gif-sur-Yvette Cedex, France § Lab Genet Biophys Plantes, Aix-Marseille Université, Marseille, F-13009, France ▽ Commissariat à l’Energie Atomique (CEA), DSV, IBEB, Marseille, F-13009, France ¶ Institut Charles Gerhardt, MACS, UMR 5253 CNRS-ENSCM-UM1-UM2, 8, rue de l’Ecole Normale, 34296 Montpellier Cedex 5, France ‡

ABSTRACT: New information on a protein’s structure, intraand intermolecular hydrogen bonds, or metal−ligand bond properties can be unraveled in the far-infrared (far-IR) terahertzdomain (600−3 cm−1 or 18−0.1 THz). In this study, we compare the performances of thermal sources with synchrotron far-IR to record reaction-induced Fourier transform infrared (FT-IR) difference signals with proteins in solution. Using the model protein Cu−azurin placed in a short path length electrochemical cell adapted for transmission spectroscopy in vacuum-purged optics, we show that minute spectral shifts induced by metal isotope labeling or temperature changes are detected using the far-IR beamline AILES of the synchrotron SOLEIL. On one hand, these data allow us to identify modes involving Cu−ligand vibrations and pave the way for the analysis of metal sites or metal redox states of proteins not amenable to resonance Raman spectroscopy. On another hand, small band shifts or changes in band intensity upon temperature modifications show that far-IR difference spectroscopy allows one to extract from a complex background hydrogenbonding signatures directly relevant to the protein function. For Cu−azurin, a temperature-sensitive IR mode involving Cu(II)− His vibrations points to the role of a hydrogen bond between a Cu histidine ligand and the water solvent in tuning the Cu(II)− histidine bond properties. Furthermore, these experimental data support the possible role of a His117−water interaction in electron-transfer activity of Cu−azurin proposed by theoretical studies.

C

sources are used for these studies, while increasing interest in the utilization of SR sources is associated with SR-based FT-IR microscopy, which provides molecular imaging at subcellular level in biological tissues5−9 or real-time monitoring of changes in cell’s or microorganisms’ physiology.5,10 Analysis of proteins or biological molecules in the far-IR terahertzdomain (600−3 cm−1 or 18−0.1 THz) is also of great interest, since it provides new information on biomolecular structures, dynamics, and properties of intraand intermolecular hydrogen bonds.11−13 Bending vibrational modes of amino acids as well as metal−ligand vibrations also absorb in the far-IR, which is particularly appealing for probing metal active sites in metalloproteins.14−19

hemical information in life-related systems is collected at the molecular level using spectroscopy. Synchrotron facilities provide stable light sources, broad bands, and brightness and give access to the whole hard X-rays to farinfrared (far-IR)terahertz (THz)spectral domain. These performances make the synchrotron radiation (SR) sources unique to study small, diluted, or strong absorbing samples leading to incomparable signal-to-noise ratio (i.e., data quality) and then spectral resolution. Mid-IR (5000−600 cm−1) spectroscopy has been extensively used to analyze the structural properties of proteins.1,2 Structural details of protein active sites and proton-transfer reactions associated with enzymatic activities are also monitored with a high sensitivity using reaction-induced Fourier transform infrared (FT-IR) difference spectroscopy.3,4 This approach circumvents bulk absorption of the water solvent and of residues not involved in the targeted reaction. Thermal © 2013 American Chemical Society

Received: December 4, 2012 Accepted: January 28, 2013 Published: January 29, 2013 2891

dx.doi.org/10.1021/ac303511g | Anal. Chem. 2013, 85, 2891−2898

Analytical Chemistry

Article

Figure 1. Technical drawings of the electrochemical cell. Two 1° wedged CVD diamond windows (Φ = 10 mm, 300 μm mean width) (a) are fixed on two Delrin rings (b) that accommodate two Viton O-rings (c). The Delrin rings adapt into an epoxy body (d), and the cell chamber is sealed by squeezing the two faces of a thermostated copper holder (e). The working electrode (f) is a 4−6 μm thick chemically modified gold grid deposited on one of the windows (g). The counter electrode (h) is a thin platinum foil deposited at the window periphery, and the reference electrode (i) consists in a silver (Ag) wire covered by AgCl (j) and immersed in a 3 M KCl solution (k) (ref 37). The electrode is screwed in the epoxy body, and the tightness is ensured by an O-ring (c). It is in electrical contact with the cell chamber via a cotton linter (l). The sample is deposited on one of the windows before placing the second window and adjusting the path length. A hole (m) in the epoxy body allows filling the space at the window’s periphery with buffer to achieve electrical contact between the sample and the counter and reference electrodes. The working and counter electrodes are connected to steel nails (n) plugged in nylon screws (o) via 1 mm thick platinum (Pt) wires (p) passing through the epoxy body. All connections are tightened using silicone sealants (q) and/or O-rings (c).

All of this enables us to probe metal site properties and the role of protein−solvent interactions in the turnover of redox metalloproteins in solution.

Far-IR absorption spectra of proteins in aqueous solution are dominated by the strong absorption of water,20,21 and FT-IR difference spectroscopy is needed to identify far-IR modes associated with metalloprotein active sites. Yet, only a few studies were reported in the literature, concerning light-induced reactions on partially dehydrated photosynthetic proteins or electrochemically induced redox transitions in metalloproteins and related models.16−18,21−27 This scarceness is related to the weakness of difference signals, typically of the order of 10−5 to 10−3 AU. This approach also faces technical difficulties caused by extensive water and water vapor absorption, together with limitations in the choice of optical materials, source brilliance, and detectors sensitivity. Finally, the data interpretation is not straightforward, since many bands can overlap in the difference spectra and since the far-IR modes are often delocalized on a large number of atoms in the far-IR region. In the present study, we compare the performances of setups involving purged or vacuum optics and thermal Globar or SR IR sources for electrochemically induced far-IR FT-IR difference spectroscopy. Using the model protein Cu−azurin and a tight thin path length electrochemical cell adapted for transmission spectroscopy in vacuum-purged optics, we show that minute spectral shifts induced by metal isotope labeling can be detected using the SR far-IR beamline AILES of the synchrotron SOLEIL.28,29 Taking advantage of the high spectral quality obtained on the beamline below 300 cm−1, we recorded unprecedented temperature effects in electrochemically induced FT-IR difference spectra in the low-frequency region (≈200 cm−1), where libration modes of water as well as intraor inter-hydrogen-bonding modes are expected to contribute.

■

EXPERIMENTAL METHODS

Preparation of the Protein Samples. Escherichia coli BL21 (DE3) strain (purchased from Invitrogen) was transformed with the pGK22(1) plasmid containing the Pseudomonas aeruginosa Cu−azurin azu gene.30,31 For heterologous expression, cells were grown overnight at 37 °C under aerobic conditions in LB medium supplemented with 100 μg/mL ampicillin. Cu−azurin was purified as previously described32 using HiTrap SP FF (GE Healthcare) and HiTrap DEAE sepharose FF (GE Healthcare) columns instead of Whatman CM23 and DE52 columns, respectively. Samples of 63Cu−azurin or 65Cu−azurin were prepared following literature methods33,34 with some modifications. To prepare apoazurin (i.e., azurin deprived of Cu), Cu−azurin (2 mg·mL−1) was incubated for 1 h at room temperature under stirring with 0.5 M KCN in 20 mM Tris−HCl pH 7. This was followed by two dialysis steps against 20 mM Tris−HCl pH 7 and one against 20 mM acetate pH 4, at 4 °C. 63Cu−azurin or 65 Cu−azurin was then obtained by incubation of apoazurin with 63 CuCl2 or 65CuCl2 (prepared from 63CuO or 65CuO purchased from Euriso-top, France) during 30 min at room temperature under moderate shaking at a Cu/apoazurin ratio of five. Cu in excess was removed by dialysis against 20 mM Tris−HCl pH 7 at 4 °C. 2892

dx.doi.org/10.1021/ac303511g | Anal. Chem. 2013, 85, 2891−2898

Analytical Chemistry

Article

in the 450−50 cm−1 region using either the dry-air-purged or the evacuated spectrometers. Notice that the dry air purging flux may also cause vibrations of optical materials, notably the 6 μm thick Mylar beam splitter used to access the far-IR domain. In order to combine electrochemical experiments with the spectrometer under vacuum (pressure of 10−5 mbar), we developed a vacuum-tight electrochemical cell, on the basis of that described by Moss et al.37 with modifications to access the far-IR domain.21 Drawings of the cell are presented in Figure 1. Using this cell, we recorded FT-IR difference spectra in the 450−50 cm−1 domain using the model protein Cu−azurin (see below). Reduced-minus-oxidized FT-IR difference spectra obtained at 4 cm−1 resolution using the Globar source on a dry-airpurged Bruker 66SX spectrometer and on the vacuum-purged IFS125 Bruker spectrometer of the AILES beamline are displayed in Figure 2, parts a and b. These spectra resulted from averaging over the same number of scans. It is noteworthy that the spectra were collected in 10.2 h at the laboratory and in 4.2 h at the beamline because of the greater mobile mirror

Electrochemistry. The electrochemical cell is presented in Figure 1. Two 1° wedged chemical vapor deposition (CVD) diamond windows were purchased from Crystran Ltd. (U.K.). The working gold grid purchased from Euromip (France) was surface-modified by dipping it for 10 min into a 5 mM pyrimide-3-carboxyaldehyde thiosemicarbazone (PATS-3, Lancaster) solution heated to 80−90 °C. The following redox mediators (each at 60 μM final concentration) were used to accelerate the electrochemical reaction: ferricyanide (Em = 436 mV vs normal hydrogen electrode, NHE), tetramethyl-pphenylenediamine (Em = 270 mV vs NHE), and phenazine ethosulfate (Em = 55 mV vs NHE). Electrochemically induced FT-IR difference spectroscopy experiments were performed with 10 mM Cu−azurin in 50 mM Tris−HCl 100 mM KCl pH 8.5. An amount of 15 μL of sample was then deposited between the two CVD windows and the cell path length was adjusted manually until the IR absorption of the sample at 1643 cm−1 (δ(O−H)H2O contribution35) was less than 0.9 absorbance units (AU). The electrochemical cell was thermostatted at 2, 8, or 36 °C within 0.1 °C with a water circulation system using a PT100 sensor connected to a controller (Huber cc 805). A potentiostat (EG&G 362, Princeton Applied Research or Microstat, Sycopel Scientific Ltd.) triggered by the FT-IR spectrometer was used to apply the oxidizing (560 mV vs NHE) and reducing (−40 mV vs NHE) potentials to the electrochemical cell. After the change in redox potential, a 5 min delay was awaited prior to the spectral measurements, for ensuring complete oxidation or reduction of Cu−azurin (Em = 293 mV vs NHE at pH 836). FT-IR Spectroscopy Measurements. FT-IR spectra in the 450−50 cm−1 range were measured at 4 cm−1 resolution using a dry-air-purged spectrometer (Bruker 66 SX) or the vacuumevacuated (10−5 mbar) Bruker IFS125 spectrometer of the farIR terahertz beamline AILES connected to the synchrotron source SOLEIL. The two spectrometers are outfitted with 6 μm mylar beam splitters and Si bolometer detectors (Infrared Laboratories, U.S.A.) with a 690−20 cm−1 cold filter and Globar conventional light sources (GCS). Each single spectrum resulted from the averaging of 300 or 500 scans recorded at 10 or 40 kHz, respectively, with the Bruker 66 SX and the IFS125 Bruker spectrometers. For each sample, results from 10 to 20 successive independent redox cycles were averaged. Spectra were recorded at 8 °C except otherwise mentioned. Noise Determination. The noise root mean square (RMS) was calculated in the 260−200, 187−149, and 100−50 cm−1 domains, using the command Signal to Noise ratio of the OPUS 7.0 Spectroscopy software. A noise RMS was calculated for each redox cycle allowing us to evaluate the noise evolution with scan numbers and to compare the noise RMS level after a defined acquisition time (same number of scans) for each setup configuration.

■

Figure 2. Influence of vacuum-purged optics and SR source on far-IR FT-IR difference spectra. Reduced-minus-oxidized FT-IR difference spectra recorded on Cu−azurin samples at 4 cm−1 resolution, with GCS sources and a dry-air-purged Bruker 66SX spectrometer (a) or the Bruker IFS125 spectrometer of the AILES beamline (b) and with the SR source and the Bruker IFS125 spectrometer (c). Each spectrum corresponds to a total of 30 000 scans. Noise RMS as a function of the number of averaged scans in the 260−200 (d), 187−149 (e), and 100−50 cm−1 (f) spectral ranges. In panels d−f, squares, triangles, and circles correspond to configurations used to record spectra shown in panels a−c, respectively. Measurements and noise calculation conditions are detailed in the Experimental Methods section (AU, absorbance unit).

RESULTS AND DISCUSSION Electrochemically Induced Far-IR FT-IR Difference Spectroscopy Using Vacuum-Purged Optics and the Synchrotron Source. In previous experiments performed with a dry-air-purged Bruker 66SX spectrometer, we observed that water vapor contributes significantly below 350 cm−121 and that averaging spectra over several electrochemical cycles (typically 20−30 cycles) is needed to reduce the fluctuation of signal caused by slight variations of the hydration level in the spectrometer. In the present study, we compare the data quality 2893

dx.doi.org/10.1021/ac303511g | Anal. Chem. 2013, 85, 2891−2898

Analytical Chemistry

Article

speed at the beamline (40 vs 10 kHz). Positive and negative bands are perfectly reproducible in these spectra, and the signalto-noise ratio is equivalent above ≈350 cm−1, while it is significantly enhanced in the spectrum recorded with the vacuum-purged optics below 350 cm−1. The reduced-minusoxidized FT-IR difference spectrum recorded using the SR beamline shows further improved spectral quality below 300 cm−1 (Figure 2c). Parts d−f of Figure 2 show the evolution of the spectral noise RMS as a function of the number of averaged scans below 260 cm−1. Between 260 and 149 cm−1 (Figure 2, parts d and e), the noise level of the spectra recorded with the dry-air-purged spectrometer is significantly larger than the one obtained with vacuum-purged optics, and it decreases slowly with the number of accumulated scans. In contrast, noise smaller than 3 × 10−5 AU is reached in all the ranges for data recorded using the evacuated spectrometer. The lower signal-to-noise ratio at low frequency is caused by the strong absorption of water vapor in this far-IR region, which cannot be totally compensated in dryair purge measurements. In this spectral range, the use of the SR source significantly increases the spectral quality for short accumulation times so that high-quality difference spectra are obtained with a very small number of redox cycles. Below 100 cm−1, the Globar emissivity decreases.38 Consequently, the noise reaches higher thresholds for both dry-air-purged and evacuated spectrometers (15 × 10−5 and 7.5 × 10−5 AU, respectively, Figure 2f). Moreover, longer data accumulation with the dry-air-purged spectrometer has a limited effect on the noise level below 100 cm−1. This noise level is too important to obtain reliable band assignment below 100 cm−1 in the FT-IR difference spectra. In contrast, the use of the SR source increases the spectral quality by a factor of 4 in this spectral domain (Figure 2f). This is notably due to the brilliance of the SR source, which is as much as 3 orders of magnitude greater than that of GCS below 100 cm−1.38 These results show that the combination of the SR IR source and evacuated optics is advantageous over other setups to record well-defined FT-IR difference spectra in short time at 4 cm−1 resolution below 450 cm−1, making this setup optimal for the far-IR analysis of time-resolved reactions in proteins. Furthermore, only the SR radiation based setup provides the spectral quality necessary for the reliable analysis of the 100−50 cm−1 THz domain by means of FT-IR difference spectroscopy. Signal-to-Noise Ratio and Spectral Resolution. Far-IR spectroscopy for probing metal−ligand bond properties benefits strongly from metal isotope labeling for band assignments. Since very small band shifts are expected upon such labeling experiments (see below), one aim of this study was to define the best adapted spectral resolution for detecting these band shifts. Figure 3a shows reduced-minus-oxidized FT-IR difference spectra recorded with Cu−azurin in the 450−50 cm−1 domain at 4, 2, or 1 cm−1 resolution, using the SR-FT-IR setup. The spectra result from the adding of the same number of scans. Similar band profiles, bandwidths, and peak maxima are observed in the spectra recorded at 4 and 2 cm−1 resolution. As shown in Figure 3b−d, the noise level in the two spectra is almost similar in the 260−200 cm−1 range. Below 200 cm−1, the noise level decreases more slowly with data accumulation in the spectrum recorded at 2 cm−1 than at 4 cm−1, but remains sufficient (below 2 × 10−5 AU) for a reliable measurement. In contrast, the noise level appears significantly higher in the spectrum recorded at 1 cm−1 resolution especially at lowest

Figure 3. Effect of the spectral resolution on the far-IR FT-IR difference spectra. Reduced-minus-oxidized FT-IR difference spectra recorded on 10 mM Cu−azurin samples at 4, 2, and 1 cm−1 resolution in the 450−50 cm−1 IR domain (a) using the SR source and the Bruker IFS125 spectrometer of the AILES beamline. The spectra correspond to accumulation times of 6.2, 7.2, and 11.4 h for the 4, 2, and 1 cm−1 resolution, respectively. The noise RMS was calculated as a function of acquisition time in the 260−200 (b), 187−149 (c), and 100−50 cm−1 (d) spectral ranges. In panels b−d, circles, triangles, and squares represent noise evolution obtained at 4, 2, and 1 cm−1 resolution, respectively. See the Experimental Methods section for measurements and noise calculation conditions (AU, absorbance unit).

wavenumbers and decreases slowly with measurement time (Figure 3b−d). This noise observed in the whole spectral range (450−50 cm−1, Figure 3a) is due to the appearance of fringes caused by multiple reflections in the CVD diamond windows. Indeed, although the use of 1° wedged windows significantly reduces this fringing, it is not possible to avoid it completely. This artifact modifies the band profiles and the position of the peak maxima, as well-exemplified for the negative and positive bands at 303 and 276 cm−1 (Figure 3a). In short, good spectral quality is obtained at 2 cm−1 resolution in the whole 450−50 cm−1 range with a reasonable acquisition time. In addition, data recorded at 4 and 2 cm−1 show that the bandwidths are not modified by the spectral resolution. A distribution of slightly different conformations of the Cu active site in the solution of Cu−azurin, i.e., a distribution of slightly different interactions formed by the chemical groups with their environment, probably accounts for this extended width. 2894

dx.doi.org/10.1021/ac303511g | Anal. Chem. 2013, 85, 2891−2898

Analytical Chemistry

Article

By increasing the spectral resolution to 2 cm−1, we reduce the uncertainty in the band position (Δν), which is proportional to the square root of the number of data points included in the bandwidth and also depends on the line shape of the band.39 This complex relation can be approximated as follows: Δν ≈

ΔW 2(S/N)

with ΔW the bandwidth at half-maximum and (S/N) the signal-to-noise ratio. For the band at 407 cm−1 we calculated a width at halfmaximum of 8.7 cm−1 which, combined with the high signal-tonoise ratio of ∼50 obtained with the synchrotron source at 2 cm−1 resolution, results in a precision of the band position better than 0.1 cm−1. The peak maxima of the other bands studied in detail below are defined with an equivalent precision. In the following, spectra were thus recorded at 2 cm−1 resolution to benefit from this high precision in band position. Identification of Metal−Ligand IR Modes Using Metal Isotope Labeling. IR modes involving Cu−ligand vibrations in Cu−azurin are expected to contribute below 450 cm−1.40 Only vibrations of chemical groups selectively perturbed by the Cu redox switch contribute in the difference spectra displayed in Figures 2 and 3. Well-defined bands are observed for the Cu(II) state (negative bands) at 407, 345, 303, 258, 190, and 114 cm−1, and for the Cu(I) state (positive bands) at 447, 311, 276, 225, and 145 cm−1. To identify IR modes involving Cu−ligand vibrations, we compared difference spectra recorded with 63Cu−azurin and 65 Cu−azurin samples at 2 cm−1 spectral resolution with the SRFT-IR setup (Figure 4). Upon 63Cu exchange by 65Cu, the spectra are almost identical (Figure 4a) except the two negative bands at 407 and 303 cm−1 (Cu(II) state) that are downshifted by 0.8 and 0.6 cm−1, respectively (Figure 4, parts b and d). In contrast, the negative band at 345 cm−1 remains unchanged upon the substitution (Figure 4c). We can thus conclude that the bands at 407 and 303 cm−1 correspond to stretching modes involving Cu(II). No effect of Cu labeling is clearly detected on the positive bands corresponding to the Cu(I) state. Although these bands are less intense than the negative ones, we can rule out any large downshift on these bands upon 63Cu/65Cu exchange. The Cu atom of Cu−azurin adopts a bipyramidal trigonal coordination27,41 with strong interactions with the equatorial ligands provided by a cysteine sulfur (Cys112) and two histidinyl imidazole nitrogens (His46 and His117, Figure 5). Two weak axial pseudoligands are provided by a methionine thioether (Met121) and the carbonyl oxygen of a glycine (Gly45).41−43 Cu−azurin is involved in electron-transfer reactions, and structural changes associated to the Cu redox switch are directly relevant to its function.43,44 This protein being a model for the blue or type I redox Cu centers was extensively studied by numerous spectroscopic techniques,33,36,42−44 including resonance Raman (RR) spectroscopy.33,43 This technique is restricted to the oxidized Cu(II) state, which gives rise to a characteristic absorption band at 627 nm in the visible range, while the Cu(I) state presents no absorption in this spectral domain.45 The RR experiments on Cu−azurin revealed that four bands near 400 cm−1 were shifted by −0.2 to −1 cm−1 upon 65 Cu/63Cu exchange.33 These bands, showing smaller shifts

Figure 4. Effect of 63Cu−azurin and 65Cu−azurin isotopic labeling on the FT-IR difference spectra. (a) Superimposition of reduced-minusoxidized FT-IR difference spectra recorded at pH 8.5 on 10 mM of 63 Cu−azurin (blue) and 65Cu−azurin (red) samples at 2 cm−1 resolution, using the SR source and the Bruker IFS125 spectrometer. Spectra correspond to a total of 33 000 and 29 000 scans for 63Cu− azurin and 65Cu−azurin, respectively. Zoom of the 412−402 (b), 353−337 (c), and 308−295 cm−1 (d) regions.

Figure 5. Cu−azurin active site (PDB 4AZU, ref 41) drawn using Accelrys DS Visualizer program.

than that expected for a pure Cu−S(Cys) stretching (ν) mode (greater than 2 cm−146), were assigned to a coupling of the ν(Cu−SCys) stretching mode with bending modes of the cysteine side chain.46−48 The band observed at 407.3 cm−1 in the far-IR FT-IR difference spectrum recorded with 63Cu− azurin is in line with the position of the intensity-weighted average Cu−S(Cys) stretching frequencies of the four RR modes.34 It is assigned to the main IR vibrational band involving the ν(Cu−SCys) mode. The small but significant 2895

dx.doi.org/10.1021/ac303511g | Anal. Chem. 2013, 85, 2891−2898

Analytical Chemistry

Article

downshift of this IR band upon 63Cu/65Cu exchange (−0.8 cm−1) confirms that the ν(Cu−SCys) mode is strongly coupled with deformation modes of amino acids. A weak band at 307 cm−1 with no clear effect of 63Cu/65Cu labeling was reported in RR studies of azurin.33 It was proposed to account for the νas(Cu−(NHis)2) on the basis of Cu/Ni substitution.34,48 In contrast to RR, asymmetric stretching vibrations contribute strongly in far-IR FT-IR. The clear downshift by 0.6 cm−1 of the far-IR band at 303.5 cm−1 upon 63 Cu/65Cu exchange shows that stretching modes involving Cu(II) contribute at this frequency and correspond most likely to the νas(Cu−(NHis)2) IR mode coupled with bending modes of the Cu ligands. These first far-IR data on a type I Cu center illustrate the potential of spectroscopy in this domain for providing clear fingerprints of protein metal centers. The high-resolution spectra obtained using SR sources allow the identification of metal−ligand vibrational modes using shifts induced by metal labeling, a strategy that overcomes metal substitution studies as it does not change the metal center properties.34,48 Moreover, in the present sample, the far-IR data provide the first vibrational data on the reduced Cu(I) center not amenable using RR spectroscopy. These early spectra will be supplemented by recording a series of labeled derivatives, such as H/2H-exchanged sample, and by comparing experimental data with normal mode predictions using quantum chemical calculations. Such comprehensive analysis of the farIR spectra will contribute to a better understanding of metal site geometries and the physical properties of proteins. Probing Hydrogen Bonding by the Detection of Temperature-Sensitive Signatures in the Far-IR Difference Spectra. Protein absorption spectra below 300 cm−1 are dominated by a large number of delocalized modes as well as by contributions from vibrational bands due to intra- and intermolecular ν(H···O) or ν(H···N) hydrogen bonds and contributions from the water solvent.11,20,49−55 Properties of hydrogen bonds in proteins, and notably those involving the water solvent, are highly sensitive to the temperature.56,57 The thermal dependence of far-IR absorption spectra of proteins or peptides has been studied in order to specifically probe modes corresponding to these hydrogen-bonding interactions.51,53,54 However, the hydrogen-bonding signature is complex in absorption spectra, and it is not easily accessible for proteins in solution. In this study, we analyzed the effect of temperature on reaction-induced far-IR difference spectra to test if the present experiment allows the investigation of hydrogenbonding properties directly relevant to the protein function. Figure 6 shows a superimposition of reduced-minus-oxidized difference spectra recorded at 2 and 36 °C with Cu−azurin in H2O buffer. The spectra appear similar, except at 225−150 cm−1, where the amplitude of the negative broad signal is increased at 36 °C. In addition, the negative band at 303.7 cm−1 in the spectrum recorded at 2 °C is significantly downshifted to 302.7 cm−1 at 36 °C. The negative band at 225−150 cm−1 is associated to the Cu(II) state. It is close to the intermolecular connectivity band, arising from the longitudinal motion of the hydrogen atom along the hydrogen bond axis (i.e., hydrogen bond stretching). Interpretation of this band is not straightforward, since several factors such as the number of hydrogen bonds but also the polarization and the dynamics of the vibration can influence the absorption intensity in the far-IR domain (ref 55 and references therein). The far-IR difference spectra show, however, that the

Figure 6. Temperature dependence of the FT-IR difference spectra. Superimposition of reduced-minus-oxidized FT-IR difference spectra recorded at 36 °C (red) and 2 °C (blue) on Cu−azurin samples at 2 cm−1 resolution (a) using the SR source and the Bruker IFS125 spectrometer. The spectra recorded at 36 and 2 °C correspond to a total of 23 000 and 26 000 scans, respectively. The main temperature effects are zoomed in panels b and c.

Cu redox switch involves changes with temperature suggesting a dependence on hydrogen bonding. The band at 303 cm−1 has been shown to contain significant contribution from the Cu(II)−ligand stretching mode, most likely ν(Cu−(NHis)2) vibration (see above). Interestingly, the histidine Cu ligand His117 is hydrogen-bonded to a water solvent molecule, in the crystallographic structure of Cu−azurin (Figure 5),41 as well as in that of Cu−azurin dimers obtained by chemical cross-link.58 A weakening of this hydrogen bond at 36 °C could explain the temperature dependence of the band frequency at ≈303 cm−1. The frequency downshift observed upon temperature increase is in line with a decreased electronegativity of the imidazole ligand due to a weakening of the imidazole NH···OH2 intermolecular bond. A weakening of hydrogen bonds formed with the solvent is expected with increasing temperatures, as reported between the peptide backbone of α helix structures and the water solvent.56 The experimental evidence for the role of hydrogen bonding in monitoring the properties of a histidine Cu ligand is of high interest, since it has been proposed that the hydrogen-bonding interaction between His117 and a water molecule modulates the electron-transfer kinetics between Cu−azurin monomers and hence between Cu−azurin and its redox partners in solution.43,58−63 2896

dx.doi.org/10.1021/ac303511g | Anal. Chem. 2013, 85, 2891−2898

Analytical Chemistry

Article

(10) Goff, K. L.; Quaroni, L.; Wilson, K. E. Analyst 2009, 134, 2216− 2219. (11) Plusquellic, D. F.; Siegrist, K.; Heilweil, E. J.; Esenturk, O. ChemPhysChem 2007, 8, 2412−2431. (12) Ueno, Y.; Ajito, K. Anal. Sci. 2008, 24, 185−192. (13) Smye, S. W.; Chamberlain, J. M.; Fitzgerald, A. J.; Berry, E. Phys. Med. Biol. 2001, 46, R101−R112. (14) Matei, A.; Drichko, N.; Gompf, B.; Dressel, M. Chem. Phys. 2005, 316, 61−71. (15) Trivella, A.; El Khoury, Y.; Gaillard, T.; Stote, R. H.; Merino, N.; Blanco, F. J.; Hellwig, P. AIP Conf. Proc. 2010, 1214, 3−6. (16) Marboutin, L.; Desbois, A.; Berthomieu, C. J. Phys. Chem. B 2009, 113, 4492−4499. (17) Marboutin, L.; Petitjean, H.; Xerri, B.; Vita, N.; Dupeyrat, F.; Flament, J. P.; Berthomieu, D.; Berthomieu, C. Angew. Chem., Int. Ed. 2011, 50, 8062−8066. (18) El Khoury, Y.; Hellwig, P. ChemPhysChem 2011, 12, 2669− 2674. (19) Jalilehvand, F.; Mah, V.; Leung, B. O.; Janos, M.; Bernard, G. M.; Laszlo, H. Inorg. Chem. 2009, 48, 4219−4230. (20) Brubach, J. B.; Mermet, A.; Filabozzi, A.; Gerschel, A.; Roy, P. J. Chem. Phys. 2005, 122, 184509. (21) Berthomieu, C.; Marboutin, L.; Dupeyrat, F.; Bouyer, P. Biopolymers 2006, 82, 363−367. (22) Chu, H. A.; Debus, R. J.; Babcock, G. T. Biochemistry 2001, 40, 2312−2316. (23) Kimura, Y.; Mizusawa, N.; Ishii, A.; Nakazawa, S.; Ono, T. J. Biol. Chem. 2005, 280, 37895−37900. (24) Kimura, Y.; Mizusawa, N.; Ishii, A.; Yamanari, T.; Ono, T. A. Biochemistry 2003, 42, 13170−13177. (25) Yamanari, T.; Kimura, Y.; Mizusawa, N.; Ishii, A.; Ono, T. A. Biochemistry 2004, 43, 7479−7490. (26) Chu, H. A.; Sackett, H.; Babcock, G. T. Biochemistry 2000, 39, 14371−14376. (27) Adman, E. T.; Stenkamp, R. E.; Sieker, L. C.; Jensen, L. H. J. Mol. Biol. 1978, 123, 35−47. (28) Roy, P.; Brubach, J.; Manceron, L.; Rouzieres, M.; Pirali, O.; Creff, G.; Kwabia-Tchana, F.; Peng, W. In 2010 35th International Conference on Infrared Millimeter and Terahertz Waves (IRMMW-THz); IEEE Conference Publications: Rome, 2010; pp 1−2. (29) Roy, P.; Rouzières, M.; Qi, Z.; Chubar, O. Infrared Phys. Technol. 2006, 49, 139−146. (30) Arvidsson, R. H.; Nordling, M.; Lundberg, L. G. Eur. J. Biochem. 1989, 179, 195−200. (31) Karlsson, B. G.; Pascher, T.; Nordling, M.; Arvidsson, R. H.; Lundberg, L. G. FEBS Lett. 1989, 246, 211−217. (32) van de Kamp, M.; Hali, F. C.; Rosato, N.; Agro, A. F.; Canters, G. W. Biochim. Biophys. Acta 1990, 1019, 283−292. (33) Blair, F. G.; Campbell, G. W.; Cho, W. K.; English, A. M.; Fry, H. A.; Lum, V.; Norton, K. A.; Schoonover, J. R.; Chan, S. I. J. Am. Chem. Soc. 1985, 107, 5755−5766. (34) Fitzpatrick, M. B.; Czernuszewicz, R. S. JBIC, J. Biol. Inorg. Chem. 2009, 14, 611−620. (35) Venyaminov, S.; Kalnin, N. N. Biopolymers 1990, 30, 1243− 1257. (36) Pascher, T.; Karlsson, B. G.; Nordling, M.; Malmstrom, B. G.; Vanngard, T. Eur. J. Biochem. 1993, 212, 289−296. (37) Moss, D.; Nabedryk, E.; Breton, J.; Mantele, W. Eur. J. Biochem. 1990, 187, 565−572. (38) Griffiths, P. R.; Homes, C. C. Instrumentation for Far-Infrared Spectroscopy. Handbook of Vibrational Spectroscopy [Online]; John Wiley & Sons, Ltd., 2006. http://onlinelibrary.wiley.com/doi/10. 1002/0470027320.s0207/abstract (accessed February 14, 2013). (39) Davis, S. P.; Abrams, M. C.; Brault, J. J. W. Fourier Transform Spectrometry; Academic Press: San Diego, CA, 2001. (40) Nakamoto, K. Infrared and Raman Spectra of Inorganic and Coordination Compounds, Part A and B, 6th ed.; Wiley: New York, 2009.

This study demonstrates that coupling far-IR difference spectroscopy with the monitoring of temperature changes is a powerful tool for analyzing hydrogen bonding involving the solvent. Not only does it allow extracting the hydrogen bond signature from a complex background, but it also enables us to detect hydrogen bonding that tunes the properties of the second-sphere environment of the protein active site. Consequently, this method will be useful to study the thermodynamics and the electrochemical properties of metal cofactors in metalloenzymes64−66 directly relevant to the protein function.

■

CONCLUSIONS In this work, we have demonstrated the gains induced by using both vacuum-purged optics and SR light source for exploring far-IR properties of proteins in solution. Using an adapted electrochemical cell, we optimized the experimental conditions to detect very low intensity IR difference bands corresponding to modes specifically disturbed by the Cu redox switch in Cu− azurin, as well as minute band shifts or intensity changes induced by specific metal isotopic labeling or temperature changes. These data will pave the way for probing metal interactions in metalloproteins as well as the hydrogen bonding directly associated with the protein function. The investigation of redox biological systems in the far-IR domain will be a useful complement to X-ray crystallography or NMR studies for probing small, diluted, strong absorbing, or noncrystallizable samples or samples for which structural data are only available for a particular redox state.

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected] (J.-B.B.); [email protected] (C.B.). Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We acknowledge Professor G. W. Canters for giving us the plasmid containing the azu gene coding for Pseudomonas aeruginosa Cu−azurin. This work was funded in part by the French Agence Nationale de la Recherche (Grant No. ANR-08PCVI-0011).

■

REFERENCES

(1) Byler, D. M.; Susi, H. Biopolymers 1986, 25, 469−487. (2) Barth, A. Biochim. Biophys. Acta, Bioenerg. 2007, 1767, 1073− 1101. (3) Siebert, F.; Hildebrandt, P. Vibrational Spectroscopy in Life Science; Wiley-VCH Verlag GmbH & Co: Weinheim, Germany, 2008. (4) Berthomieu, C.; Hienerwadel, R. Photosynth. Res. 2009, 101, 157−170. (5) Chen, L.; Holman, H. Y.; Hao, Z.; Bechtel, H. A.; Martin, M. C.; Wu, C.; Chu, S. Anal. Chem. 2012, 84, 4118−4125. (6) Dumas, P.; Sockalingum, G. D.; Sule-Suso, J. Trends Biotechnol. 2007, 25, 40−44. (7) Marcelli, A.; Cricenti, A.; Kwiatek, W. M.; Petibois, C. Biotechnol. Adv. 2012, 30, 1390−1404. (8) Petibois, C.; Piccinini, M.; Guidi, M. C.; Marcelli, A. J. Synchrotron Radiat. 2010, 17, 1−11. (9) Nasse, M. J.; Walsh, M. J.; Mattson, E. C.; Reininger, R.; Kajdacsy-Balla, A.; Macias, V.; Bhargava, R.; Hirschmugl, C. J. Nat. Methods 2011, 8, 413−416. 2897

dx.doi.org/10.1021/ac303511g | Anal. Chem. 2013, 85, 2891−2898

Analytical Chemistry

Article

(41) Nar, H.; Messerschmidt, A.; Huber, R.; van de Kamp, M.; Canters, G. W. J. Mol. Biol. 1991, 221, 765−772. (42) Bioinorganic Chemistry of Copper; Karlin, K. D., Tyeklàr, Z., Eds.; Chapman & Hall: New York, 1993. (43) Kolczak, U.; Dennison, C.; Messerschmidt, A.; Canters, G. W. In Handbook of Metalloproteins [Online]; John Wiley & Sons, Ltd., 2006; pp 1170−1194. http://onlinelibrary.wiley.com/doi/10.1002/ 0470028637.met185/abstract (accessed February 14, 2013). (44) Groeneveld, C. M.; Feiters, M. C.; Hasnain, S. S.; van Rijn, J.; Reedijk, J.; Canters, G. W. Biochim. Biophys. Acta 1986, 873, 214−227. (45) Solomon, E. I. Inorg. Chem. 2006, 45, 8012−8025. (46) Nestor, L.; Larrabee, J. A.; Woolery, G.; Reinhammar, B.; Spiro, T. G. Biochemistry 1984, 23, 1084−1093. (47) Woodruff, W. H.; Norton, K. A.; Swanson, B. I.; Fry, H. A. Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 1263−1267. (48) Czernuszewicz, R. S.; Fraczkiewicz, G.; Zareba, A. A. Inorg. Chem. 2005, 44, 5745−5752. (49) Zelsmann, H. R. J. Mol. Struct. 1995, 350, 95−114. (50) Miura, N.; Yamada, H.; Moon, A. Spectrochim. Acta, Part A 2010, 77, 1048−1053. (51) El Khoury, Y.; Trivella, A.; Hellwig, P. IEEE Trans. Terahertz Sci. Technol. 2010, 3, 183−191. (52) Whitmire, S. E.; Wolpert, D.; Markelz, A. G.; Hillebrecht, J. R.; Galan, J.; Birge, R. R. Biophys. J. 2003, 85, 1269−1277. (53) El Khoury, Y.; Trivella, A.; Gross, J.; Hellwig, P. ChemPhysChem 2010, 11, 3313−3319. (54) Stehle, C. U.; Abuillan, W.; Gompf, B.; Dressel, M. J. Chem. Phys. 2012, 136, 075102. (55) Born, B.; Kim, S. J.; Ebbinghaus, S.; Gruebele, M.; Havenith, M. Faraday Discuss. 2009, 141, 161−173. (56) Brewer, S. H.; Tang, Y. F.; Vu, D. M.; Gnanakaran, S.; Raeigh, D. P.; Dyer, R. B. Biochemistry 2012, 51, 5293−5299. (57) Vanderkooi, J. M.; Dashnau, J. L.; Zelent, B. Biochim. Biophys. Acta, Proteins Proteomics 2005, 1749, 214−233. (58) van Amsterdam, I. M.; Ubbink, M.; Einsle, O.; Messerschmidt, A.; Merli, A.; Cavazzini, D.; Rossi, G. L.; Canters, G. W. Nat. Struct. Biol. 2002, 9, 48−52. (59) Gorren, A. C.; den Blaauwen, T.; Canters, G. W.; Hopper, D. J.; Duine, J. A. FEBS Lett. 1996, 381, 140−142. (60) Migliore, A.; Corni, S.; Di Felice, R.; Molinari, E. J. Phys. Chem. B 2006, 110, 23796−23800. (61) Migliore, A.; Corni, S.; Felice, R. D.; Molinari, E. J. Phys. Chem. B 2007, 111, 3774−3781. (62) Mikkelsen, K. V.; Skov, L. K.; Nar, H.; Farver, O. Proc. Natl. Acad. Sci. U.S.A. 1993, 90, 5443−5445. (63) van de Kamp, M.; Canters, G. W.; Wijmenga, S. S.; Lommen, A.; Hilbers, C. W.; Nar, H.; Messerschmidt, A.; Huber, R. Biochemistry 1992, 31, 10194−10207. (64) Battistuzzi, G.; Borsari, M.; Ranieri, A.; Sola, M. JBIC, J. Biol. Inorg. Chem. 2004, 9, 781−787. (65) Barker, K. D.; Eckermann, A. L.; Sazinsky, M. H.; Hartings, M. R.; Abajian, C.; Georganopoulou, D.; Ratner, M. A.; Rosenzweig, A. C.; Meade, T. J. Bioconjugate Chem. 2009, 20, 1930−1939. (66) Lay, P. A. J. Phys. Chem. 1986, 90, 878−885.

2898

dx.doi.org/10.1021/ac303511g | Anal. Chem. 2013, 85, 2891−2898