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Proton and electron transfer in wild-type and mutant reaction centers from Rhodobacter sphaeroides followed by rapid-scan FTIR spectroscopy
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Vibrational Spectroscopy 48 (2008) 126–134 www.elsevier.com/locate/vibspec
Proton and electron transfer in wild-type and mutant reaction centers from Rhodobacter sphaeroides followed by rapid-scan FTIR spectroscopy Alberto Mezzetti a,b,*, Winfried Leibl b,c a
Laboratoire de Spectrochimie Infrarouge et Raman UMR CNRS 8516, Universite´ de Sciences et Technologies de Lille, Bat C5, Cite´ Scientifique, 59655 Villeneuve d’Ascq, France b CEA, iBiTecS, Service de Bioe´nenerge´tique Biologie Structurale et Me´canismes (SB2SM), F-91191 Gif-sur-Yvette, France c CNRS, URA 2096, F-91191 Gif-sur-Yvette, France Received 17 July 2007; received in revised form 20 January 2008; accepted 29 January 2008 Available online 15 February 2008
Abstract Rapid-scan FTIR difference spectroscopy was used to investigate proton and electron transfer reactions in photosynthetic reaction centers from Rhodobacter sphaeroides. Experiments at different temperatures and in the presence of D2O have provided strong indication that a transient band at 1707 cm1 previously identified after both the 1st and the 2nd flashes [A. Mezzetti, W. Leibl, Eur. Biophys. J. 34 (2005) 921] is given by a transient protonation of the side chain of a Asp or Glu residue situated on the proton transfer pathway from the cytoplasm to the QB site. Experiments in D2O on a Asp-M17 ! Asn mutant reaction center, where the proton and electron transfer reactions are slowed down compared to the wild-type, showed that the kinetic isotope effect induced by H/D exchange slows down the electron transfer reaction after the 1st flash, confirming the strong coupling between proton and electron transfer. Rapid-scan FTIR experiments on Cd2+-treated reaction centers, in agreement with previous UV–vis measurements [P. Adelroth, M.L. Paddock, L.B. Sagle, G. Feher, M.Y. Okamura, Proc. Natl. Acad. Sci. U.S.A. 97 (2000) 13086], showed that upon addition of Cd2+, which inhibits proton uptake, the QAQB ! QAQB reaction is slowed. Interestingly, the transient 1707 cm1 band is not visible in the first spectrum recorded early after the flash. This strongly suggests its identification with a residue situated on the proton transfer pathway, which is perturbed upon metal cation binding. # 2008 Elsevier B.V. All rights reserved. Keywords: Rapid-scan FTIR; Rhodobacter sphaeroides; Photosynthetic reaction center; FTIR difference spectroscopy; Proton transfer; Electron transfer
1. Introduction Photosynthetic reaction centers (RCs) from purple bacteria are membrane enzymes which act as light-driven quinone reductases. Absorption of a photon entails an ultrafast charge separation between the so-called primary electron donor P (a special pair of bacteriochlorophyll-a molecules) and the primary acceptor QA, a quinone molecule strongly bound to the protein. A second quinone molecule, called QB, accepts two
* Corresponding author at: Laboratoire de Spectrochimie Infrarouge et Raman UMR CNRS 8516, Universite´ de Sciences et Technologies de Lille, Bat C5, Cite´ Scientifique, 59655 Villeneuve d’Ascq, France. Fax: +33 3 20 43 66 03. E-mail address: [email protected] (A. Mezzetti). 0924-2031/$ – see front matter # 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.vibspec.2008.01.019
electrons from QA in two consecutive photochemical events, as well as two protons from the cytoplasm. Protons are transferred to the QB site through a pathway formed by amino acid side chains and water molecules. The formed ubiquinol molecule QBH2 leaves the RC and is replaced by an oxidised quinone coming from the quinone pool present in the membrane. The ubiquinol is then reoxidised by another membrane enzyme, called the cytochrome bc1 complex. The overall effect of this reaction cycle is to move protons from the cytoplasm to the periplasm. The formed proton gradient is then used to synthesise ATP, necessary to power the metabolic reactions in the cell. Proton and electron transfer reactions, especially those involving quinone molecules, play a key role in bioenergetics [1]. Compared to other enzymes, in bacterial photosynthetic RCs the reactions can be easily triggered by short laser pulses.
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This makes photosynthetic RCs ideal systems to investigate general principles of quinone redox chemistry as well as proton and electron transfer in bioenergetic systems. The structure of the RC from Rhodobacter (Rb.) sphaeroides ˚ has been determined by X-ray crystallography with up to 2 A resolution [2–4] leading to a detailed knowledge of the positions of pigments, cofactors and amino acids, as well as on their relative distance and orientation. This makes possible to give a much deeper interpretation of data coming from spectroscopic techniques, in order to fully understand the mechanisms of the reactions taking place in the RCs. In addition, for Rb. sphaeroides site-directed mutagenesis is wellestablished. Knowledge of the structure also makes possible to change specific amino acids in key positions of the RC. Mutant RCs have indeed provided most of the recent relevant data on the working mechanism of the RC, especially concerning the proton transfer pathways (see [5,6] for recent reviews). It is now generally accepted that in isolated RCs the first electron transfer from QA to QB does not involve protonation of QB, which occurs only on the second reduction step. However, the electron transfer step is accompanied by a substoichiometric proton uptake from the cytoplasm towards the interior of the protein. In addition, driving force assay methods have demonstrated that the QAQB ! QAQB reaction is not rate-limited by an electron transfer but by another step [7]. Even though different processes have been proposed as the ratelimiting step [5,8], the implication of a proton transfer is strongly suggested by recent studies [6,9]. The second electron transfer between the two quinones leads to the formation of a QBH state in less than 1 ms. This electron transfer is coupled to, but not limited by, proton uptake from the cytoplasm [10]. Subsequently, QH is rapidly protonated and the formed QBH2 leaves the RC [11]. There is no definite agreement on the detailed mechanism of the two electron transfer reactions. Nevertheless, a dominant proton pathway from the cytoplasm to the QB site has been identified [5,6] which is active for proton transfer towards the QB site upon both electron transfer reactions. In both cases, protons are taken up by the His-H126 and His-H128 residues (Fig. 1), which in fact are part of the binding site of the proton transfer inhibitors Zn2+ and Cd2+ (Ni2+, another proton transfer inhibitors, binds at His-H126 and Asp-M17) [12]. Located between the entry point and the QB site are Asp-L210 and AspM17. These two residues play a crucial role in the transfer of both protons. Also the Asp-L213 residue is believed to be part of the proton transfer pathway. At or near Asp-L213, the pathways for the two protons diverge (see Fig. 1). Proton transfer to Glu-L212 (associated with the first electron transfer) occurs through intervening water molecules. H+ transfer directly to QB (associated to the second electron transfer) occurs through the Ser-L223 side chain. Water molecules located between the amino acid side chains are likely to act as connectors. FTIR difference spectroscopy has been largely used to investigate structure-function relationships in Rb. sphaeroides RCs and chromatophores, relying on the capability of the technique to monitor simultaneously reaction-induced changes
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Fig. 1. Superimposed on part of the crystal structure are the proton transfer pathways in Rhodobacter sphaeroides RCs. See text for further details. Figure is adapted from [36].
in both the cofactors and the proteins [13]. A large amount of data has been obtained by static FTIR difference spectroscopy leading to the precise identification of IR marker bands for specific redox states of cofactors ([14] and references therein), protein rearrangements [15,16] and protonation of amino acid side chains [17]. The availability of these marker bands has prompted a series of time-resolved IR difference spectroscopy investigations. Whereas early investigations used mainly single wavelength techniques [18,19], in more recent years timeresolved FTIR techniques have been used. After the pioneering experiments of Thibodeau et al. [15] and Burie et al. [20], Rb. sphaeroides RCs and chromatophores have been studied by rapid-scan FTIR [21–23] and step-scan FTIR [9,24]. Recently, an investigation combining both the rapid-scan and the stepscan technique has been reported [25]. It is also worth mentioning that very recently a chemometric approach to analyse time-resolved FTIR difference spectra of Rb. sphaeroides chromatophores has been proposed [26]. In this paper, we report new rapid-scan FTIR data aimed to better understand the proton and electron transfer reactions taking place in isolated Rb. sphaeroides RCs. The simultaneous investigation of the reactions in different conditions (different temperatures, different media—H2O and D2O, use of native and site-directed mutant RCs, presence of proton uptake inhibitors) provided new insight into the coupling between electron and proton transfer reactions. 2. Experimental 2.1. Sample preparation The site-directed mutation Asp-M17 ! Asn was constructed as described previously [27]. RCs from Rb. sphaeroides R26 and the mutant were isolated in 15 mM Tris–HCl pH 8, 0.025% lauryl dimethylamide-Noxide (LDAO), as described in [28]. The samples for FTIR measurements were prepared as previously described [29,30].
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Briefly, RCs were deposited on a CaF2 window and dried under argon. Before complete dryness, the RC film was re-hydrated with Tris buffer (see below). To guarantee full occupancy of the QB site, a fivefold excess of UQ6 (ubiquinone-6, Sigma) was added to the RC suspension. To ensure fast reduction of P+ external donors were added. Two different mixtures were used, following published protocols [23]: (i) ferrocyanide (250 mM) and tetramethyl-p-phenylenediamine (TMPD; 50 mM) buffered with Tris (pH 7; 100 mM) and (ii) 10 mM Na-ascorbate and 20 mM 2,3,5,6-tetramethyl-p-phenylenediamine (DAD) buffered at pH 8. The choice of the redox mediators determines the re-oxidation time of QBH2 between averaging cycles. A second CaF2 window was used to squeeze the sample, in order to yield an absorbance in the amide I region of the spectrum of 0.6–0.9 a.u. 2.2. FTIR difference spectroscopy experiments Rapid-scan measurements were performed using the experimental parameters as described previously [23]. A Bruker IFS 88 FTIR spectrometer equipped with a photoconductive MCT-A detector and with Opus software was used. Measurements were performed at 285 1 K, 281 1 K, 268 1 K and 260 1 K using a temperature-controlled N2 cryostat. The photoreactions were triggered by a saturating flash from a frequency-doubled Nd:YAG laser (7 ns, 20 mJ, Quantel, France). The experimental scheme used varied slightly according to the particular sample studied. For wild-type, untreated RCs, 40 interferograms were recorded in the dark and averaged. Then a first laser flash was fired and an interferogram was recorded between 2 ms and 27 ms to monitor the time evolution of the system after the flash. 73 ms after the 1st flash, a 2nd flash was fired (see
Fig. 2B). An interferogram was then recorded between 2 ms and 27 ms after the 2nd flash. Then four interferograms were recorded at increasing delay to monitor the process of ubiquinol formation. Subsequent interferograms were averaged in groups of five. The results from 3200 cycles (obtained on two different samples) were averaged to improve the signal-to-noise ratio. For mutant and Cd2+-treated wild-type RCs, 40 interferograms were recorded in the dark and averaged. Then a first laser flash was fired and an interferogram was recorded between 4 ms and 29 ms to monitor the time evolution of the system after the flash. Then four interferograms were recorded at increasing delay to monitor the process of ubiquinol formation (see Fig. 2A). Subsequent interferograms were averaged in groups of five. The results from 2000 cycles (obtained on two different samples) were averaged to improve the signal-to-noise ratio. Between cycles an appropriate delay time (3 min for the samples with DAD and Na-ascorbate as redox mediators; a time ranging from 30 s to 3 min, according to the temperature, for the samples with ferrocyanide and TMPD as redox mediators) allowed a complete relaxation of the system due to re-oxidation of QH2 by the external redox chemicals. The necessary relaxation time was determined by investigating the decay of the differential spectra at long times. At the end of the experiments, the stored interferograms were Fourier transformed to give the corresponding single beam spectra using the Opus software, applying the Mertz phase correction and the Blackman-Harris 3-term apodization function. Difference spectra at various times after the laser flash were calculated according to the formula DA(t) = log[S(t)/S(0)], where S(0) is the single beam spectrum obtained from the averaged interferogram recorded before the 1st flash and S(t) is the single beam spectrum at time t after the 1st or the 2nd flashes.
Fig. 2. Scheme for (A) experiments with 1 triggering flash; (B) experiments with 2 triggering flashes.
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3. Results and discussion 3.1. Rapid-scan FTIR spectra after the 1st and 2nd flashes at different temperatures Time-resolved FTIR difference spectra after one and two consecutive flashes recorded at 285 K, 268 K and 260 K are shown in Figs. 3 and 4. In these experiments ferrocyanide and TMPD were used as external donors in order to quickly reduce P+, which would give rise to IR absorption bands which are much more intense (by a factor of 5) than quinone, semiquinone and protein bands [31]. The ferrocyanide/TMPD mixture can reduce P+ more quickly than the conventional DAD/ascorbate mix used in most steady-state measurements and previously used also in rapid-scan FTIR measurements [21–23,25,26]. Moreover, ferrocyanide/TMPD ensures fast P+ reduction also at low temperature (260 K and 268 K). Although TMPD redox changes are known to give relatively intense IR signals at 1620 cm1 (negative signal), 1546 cm1 (positive signal), 1520 cm1 (negative), 1431 cm1 (positive) and 1382 cm1 (positive) [32] (which in several cases can be a serious drawback), no signals arising from TMPD+/TMPD are expected in the 1650–1750 cm1 region. Therefore, peaks in this spectral region can be safely attributed to protein and/or cofactor contributions. In Fig. 3 are reported the spectra recorded in the first scan after the 1st flash (between 2 ms and 27 ms after the flash) at the three temperatures, corresponding to QB/QB difference spectra. In fact bands characteristic of QB (positive band at 1479 cm1) of disappearing QB (negative band at 1264 cm1) and of protonation of the Glu-L212 side chain (1728 cm1) are visible at all three temperatures. Bands reflecting at least partial contributions from TMPD+/TMPD signals are visible at 1620 cm1 (negative), 1541 cm1 (positive) and 1523 cm1 (negative). These TMPD+/TMPD bands overlap with contributions from the RC; this is possibly the reason why they appear at
Fig. 3. Rapid-scan FTIR difference spectra at different temperatures recorded between 2 ms and 27 ms after the 1st flash in presence of ferrocyanide and TMPD in wild-type RCs. Bands arising – at least partially – from TMPD+/ TMPD contributions are marked by an asterisk.
Fig. 4. Rapid-scan FTIR difference spectra at different temperatures recorded between 2 ms and 27 ms after the 2nd flash in presence of ferrocyanide and TMPD in wild-type RCs. Bands arising – at least partially – from TMPD+/ TMPD contributions are marked by an asterisk.
a slightly different wavenumbers than found in electrochemically generated TMPD+/TMPD FTIR difference spectra of pure TMPD solutions [32]. Smaller TMPD+/TMPD contributions are present in our spectra but do not emerge as clear positive or negative bands as they are hidden by stronger spectral contributions from the protein and the cofactors. In addition, a positive band at 1707 cm1 is present at 260 K and 268 K, but absent at 285 K. This band has been previously observed at 281 K (using DAD and ascorbate as redox mediators) and 268 K (using ferrocyanide and TMPD) [23]. Experiments where more scans were measured after the 1st flash (data not shown; see also [23] where this experiment was carried out at 281 K) showed that this band quickly decays in intensity (it is absent in the following scans), in contrast to the QB band (1479 cm1) and the protonated Glu-L212 side chain band (1728 cm1) which are more stable. In our previous paper, this was interpreted as an indication that the 1707 cm1 band corresponds to a transient event associated with the formation of QB [23]. In Fig. 4 are shown the spectra recorded in the first scan (between 2 ms and 27 ms) after the 2nd flash at 285 K, 268 K and 260 K. They correspond to a QB(double reduced)/QB state. They all show a positive band at 1479 cm1, but with decreased intensity and increased broadness compared to the spectrum recorded immediately after the 1st flash. This band was previously reported at 281 K and attributed to undesired formation of QB in a portion of RCs [23]. Unfortunately the marker bands for QH2 and/or QBH formation overlap with the strong positive bands from TMPD+. Indications for QH2 and/or QBH formation can however be seen at 1490 cm1 (shoulder) as well as from the strong decrease in intensity of the 1479 cm1 band after the 2nd flash. Also the band at 1728 cm1 is present (but with decreased amplitude compared to the spectra recorded immediately after the 1st flash). The band at 1707 cm1 is present in the three spectra but with different intensities: it is strong at 268 K and 260 K, but weak at
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285 K. Experiments where more scans were measured after the 2nd flash (data not shown; see also [23] where this experiment was carried out at 281 K and 268 K) indicate that this band quickly decays in intensity. By comparing the spectra at the three temperatures after both the 1st and the 2nd flashes it clearly emerges that the intensity of the positive band at 1707 cm1 decreases as the temperature increases, whereas almost all other peaks are present with roughly the same intensity at all temperatures. If we focus our attention on the 285 K rapid-scan FTIR spectra, we notice that the 1707 cm1 band is absent in the first scan after the 1st flash, whereas it is present but very weak in the first scan recorded after the 2nd flash. As reported above, this band has previously been observed at 268 K and 281 K after both the 1st and the 2nd flashes and tentatively attributed to a transient protonation of a Glu or Asp side chain [23]. The absence (or very small amplitude) of this transient band at 285 K can indeed be easily explained by an effect of temperature on the reaction kinetics. It has been reported that proton transfer reactions in Rb. sphaeroides RC are slowed down as the temperature is lowered [9,21]. Therefore the decrease in intensity of the 1707 cm1 band with increasing temperature is compatible with (and supportive of) its assignment to a transiently protonated side chain of a Glu or Asp residue. In fact, it can be argued that such a transient protonation is visible in the first rapid-scan FTIR spectrum at 260 K and 268 K but becomes very weak (after the 2nd flash) or no longer visible (after the 1st flash) in the same time window (2–27 ms after the flash) at 285 K, because its decay is faster than the time resolution of the technique. We recall that rapidscan experiments at 281 K [23] (but performed using DAD/ ascorbate as redox mediators and at pH 8 instead of 7) showed a weak – but detectable – transient band at 1707 cm1 after both the 1st and the 2nd flashes. Although direct comparison with the present results is questionable due to different experimental conditions (notably the used redox mediators and the pH), it appears that the intensity of the 1707 cm1 band lies between those obtained at 285 K and those obtained at 268 K (not shown) which is consistent with the temperature influence deduced above on the 1707 cm1 band deduced above. A transient 1707 1 cm1 band has also been observed at 278 K by single wavelength time-resolved IR spectroscopy [19] and by time-resolved step-scan FTIR spectroscopy [9].1
3.2. Rapid-scan FTIR spectra on wild-type RCs in D2O: H/D exchange effects on band position In order to confirm the attribution of the 1707 cm1 band to a transiently protonated Glu or Asp side chain, we performed experiments in D2O, as in these conditions we should observe a
Fig. 5. Rapid-scan FTIR spectra recorded on wild-type RCs. From top to bottom: spectra recorded between 2 ms and 27 ms after the 1st flash in D2O at 285 K (continuous line) and H2O at 260 K (dotted line); spectra recorded between 2 ms and 27 ms after the 2nd flash in D2O at 285 K (continuous line) and H2O at 260 K (dotted line).
downshift of the band due to H/D exchange [33]. Rapid-scan FTIR difference spectra recorded in D2O after the 1st and the 2nd flashes at 285 K2 are shown in Fig. 5. Ferrocyanide and TMPD were used as external donors, as in the case of rapidscan FTIR spectra recorded in H2O (reported in Figs. 3 and 4). The first spectrum recorded after the 1st flash (Fig. 5) is indicative of QB reduction (positive band at 1479 cm1; not shown) as well as protonation of the Glu-L212 side chain (band at 1728 cm1); on the other hand the 1707 cm1 band is very weak. Interestingly, we observe a new positive band at 1717 cm1. A positive band at 1717 cm1 has been previously reported in steady-state QB/QB FTIR difference spectra of wild-type RCs and interpreted as consequence of H/D exchange
2
1
It should be noticed that, in contrast with [9,19] in the present experiments the proton transfer reaction was investigated after fast and complete reduction of the oxidised primary donor P. This eliminates disturbing spectral contributions arising from the intense P+/P bands and better reproduces the situation occurring under physiological conditions.
As explained in the following, recording of D2O spectra at 285 K is suggested by kinetic considerations. However, the rate of P+ reduction by external donors seems to be severely reduced in D2O at low temperatures. We observed that below 285 K complete P+ reduction before the beginning of the first scan after both the 1st and the 2nd flashes is no longer efficient. To obtain fast and highly reproducible reduction of P+ the temperature had to be set to 285 K.
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at the protonated side chain of Glu-L212 [17]. After the 2nd flash, a small contribution is present at 1728 cm1 whereas the band at 1707 cm1 is again very weak. It is useful to compare these data with those obtained in H2O in order to point out possible H/D exchange-induced band shifts. However, comparison with H2O spectra recorded at the same temperature is not a proper choice, because we have to take into account a probable kinetic isotope effect. Indeed, the proton transfer reaction associated to the QAQB ! QAQB electron transfer shows a kinetic isotope effect of 4.9 in wildtype Rb. sphaeroides RCs [25]. Therefore we expect that rapidscan spectra recorded at 285 K in H2O and D2O in the first scan after the 1st flash do not represent the very same situation. In other words, if we consider the 2–27 ms time window to roughly correspond to a snapshot at 14.5 ms after the flash, in the two cases different stages of the reaction are monitored, as the proton transfer is expected to proceed at a significantly higher rate in H2O compared to D2O. On the other hand, if we now consider the data obtained in H2O presented above, we have strong indications – consistent with previous observations [9,21] – for an increase of the rate of the proton transfer with the temperature. Indeed, a 20-degree difference in temperature (298 K vs. 278 K) was reported to correspond to a 3.5-fold difference in kinetics of the proton transfer reaction coupled to the QA QB ! QAQB electron transfer reaction [9]. This is consistent with the observed behaviour found in the Asp-L210 ! Asn/Asp-M17 ! Asn double mutant (compare Mezzetti et al. [21] and Paddock et al. [34]) where a 13-degree difference in temperature (294 K vs. 281 K) was found to correspond to a 4-fold difference in kinetics of the QAQB ! QAQB electron transfer reaction and of the associated proton transfer. It can therefore be argued that for the proton transfer reaction coupled to the first electron transfer between QA and QB it is more appropriate to compare H2O data at 260 K with D2O data at 285 K, as the temperature-induced deceleration of proton transfer in H2O roughly compensates the kinetic isotope effect in D2O. For the proton transfer reaction coupled to the 2nd electron transfer between QA and QB no literature data are available for a possible kinetic isotope effect, even though such an effect is likely to occur, as it normally affects proton-transfer reactions in protein [35]. On the other hand, the rapid-scan data shown above clearly indicate that the process involving the formation of the 1707 cm1 is slowed down as the temperature is lowered. Therefore the comparison between the 285 K spectrum in D2O and the 260 K rapid-scan spectrum in H2O seems proper also when spectra recorded in the first scan after the 2nd flash are compared. In Fig. 5 (dotted lines) we have reported the rapid-scan spectra after the 1st and the 2nd flashes in H2O at 260 K to allow a direct visual comparison.3
3 Intensities have been normalized to yield identical amplitudes of the 1264 cm1 band which is given by a C–O–CH3 mode of a methoxy group.
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Even though a more accurate approach would require calculation of double difference spectra (D2O minus H2O),4 some features are clearly visible from the direct comparison of the D2O and H2O spectra. The 1707 cm1 band is clearly sensitive to H/D exchange, as its intensity is considerably lower in D2O. This strengthens the identification of this band as the C O stretching of a protonable residue (Asp or Glu). One possibility is that in D2O it downshifts at 1689 cm1 (compare the spectra in H2O and D2O after both the 1st and the 2nd flashes). However, the 1689 cm1 band could also be (at least partially) given by the downshifting of the 1695 cm1 band. Another interesting feature is the intense negative band at 1687 cm1 present in spectra in H2O recorded immediately after both the 1st and the 2nd flashes. Whereas this band is absent in D2O spectra, in H2O it was found to be still present also at long delays after the 1st [21] and the 2nd flashes [23]. It has been tentatively assigned to an amide I contribution [36]. 3.3. Rapid-scan FTIR spectra on wild-type and mutant RCs in D2O: kinetic isotope effect on proton and electron transfer Comparison of time-resolved FTIR difference spectra in H2O and D2O can be used also to investigate the possible consequences of an isotope kinetic effect. In fact, IR spectroscopy can allow to investigate other possible slowing effects due to H/D exchange. We recall that in Rb. sphaeroides RCs IR marker bands are available for QA, QB, QA, QB [14], QH2 [22,23], protein rearrangements [15,16], and protonated side chain of amino acids [17]. Compared to normal UV–vis monitoring of the QA QB ! QAQB reaction, where global chromophore changes are observed, IR has the definite advantage of providing specific bands for almost any of the cofactors and molecular groups taking part in the reaction; furthermore, also the effect of H/D exchange on the position of these bands has been investigated [17]. Of course for the identification of a kinetic isotope effect we want to compare the kinetics for marker bands in H2O and D2O (possible shifts in marker band position can be easily taken into account relying on the existing literature). For this reason, the spectra to be compared must be recorded at the same temperature. As several evidences suggest that the first electron transfer between QA and QB is strongly coupled to a proton transfer reaction [5], it is possible that the observed decreased rate of 4 Calculations of double difference spectra (D2O minus H2O) is a common procedure in FTIR difference spectroscopy that can provide precise indication on H/D exchange-induced shift of bands. However this procedure requires (1) that the spectra are recorded under exactly the same conditions (except for the nature of the surrounding medium) and (2) that the spectra show an excellent signal-to-noise ratio. In time-resolved FTIR these two requirements are difficult to meet because of the intrinsic limitation in signal-to-noise ratio and because it is difficult to take properly into account the exact influence of the kinetic isotope effect (and therefore to choose the right pair of spectra for the subtraction procedure).
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proton transfer in D2O entails also a decreased rate of the electron transfer reaction. We recall that for rapid-scan spectra recorded in H2O no indication of an ongoing electron transfer is visible, even at low temperatures: in other words, the electron transfer reaction could not be resolved. For this reason we looked in the rapid-scan spectra in D2O at the spectral region where marker bands for the QAQB state lies, notably the intense positive band at 1467 cm1 [14] and the negative band at 1670 cm1 [15]. Unfortunately no traces of QAQB state can be found on the first rapid-scan spectrum recorded between 2 ms and 27 ms after the 1st flash (data not shown). Therefore, apparently in wild-type RCs at 285 K the kinetic isotope effect is incapable to slow down the electron transfer reaction between the two quinones sufficiently enough to be resolved by the rapid-scan technique. On the other hand, it is not possible to lower the temperature as in this case fast reduction of P+ is no longer efficient. We therefore looked for a mutant RC where the proton transfer coupled to the QAQB ! QAQB electron transfer reaction is already slowed down in H2O, but not enough to let QA contributions appear in the first scan. We focused our attention on an Asp-M17 ! Asn mutant which has been already investigated by UV–vis and proton uptake measurements [34], as well as static and rapid-scan FTIR [36,21]. This system is indeed ideal as in rapid-scan FTIR spectra on H2O no traces of QA were detectable in the first spectrum after the laser flash [21]. We performed some measurements on this mutant in D2O at 281 K and pH 8 (see spectra in Fig. 6), i.e. the very same conditions of temperature and pH of the published data of the Asp-M17 ! Asn mutant in H2O [21]. Also the timing of the experiment was set to be identical to the one for H2O [21], i.e. the first scan was recorded between 4 ms and 29 ms after the flash. Very interestingly, intense marker bands for the QAQB state are visible at 1467 cm1 and 1670 cm1 in the first scan after the 1st flash (Fig. 6, upper trace). Also the marker band for QB is present in the first scan (see band at 1479 cm1). Therefore, this spectrum reflects a situation in
Fig. 6. (A) Rapid-scan FTIR spectra on the Asp-M17 ! Asn mutant in D2O, T = 281 K. (B) Rapid-scan FTIR spectra on the Asp-M17 ! Asn mutant in H2O, T = 281 K; adapted from [21].
time where the electron transfer reaction is about half-way advanced. This means that the presence of D2O, combined with the 8-fold decrease caused by the mutation of the M17 amino acid [34], is capable to slow down the reaction enough to permit to observe intense QAQB state marker bands in the first scan after the laser flash. Such spectral contributions are absent in the following scans, where only bands characteristic of QB are present (see Fig. 6). Therefore, in the Asp-M17 ! Asn mutant D2O clearly affects the QAQB ! QAQB electron transfer reaction. The intrinsic mechanism of mono-electronic oxidation of QA and reduction of QB does not imply any protonation or deprotonation of the semiquinone or quinone. This means that the electron transfer reaction is rate-limited by an ‘‘external’’ step which is sensitive to H/D exchange. An interpretation would be to assume that the electron transfer is rate-limited by a proton transfer step. However, the issue on the rate-limiting step for the QAQB ! QAQB reaction is still under debate, and other ‘‘gating’’ steps sensitive to an isotopic effect are possibly involved. Nevertheless, the present results suggest a strong coupling between the QAQB ! QAQB reaction and proton transfer reactions. Unfortunately, no clear conclusions can be drawn concerning the protonation state of the Glu-L212 side chain in the first spectrum. In the following spectra, this residue is responsible for the positive bands at 1726 cm1 and 1717 cm1, as determined by steady-state FTIR difference measurements [36]; these two bands arise from the partial deuteration of the Glu-L212 residue [17,36], even though in steady-state spectra their position was slightly different (1728 cm1 and 1716 cm1). In the first spectrum (recorded between 4 ms and 29 ms after the flash), the 1726 cm1 band is overlapping with a 1735 cm1 (negative)/1727 cm1 (positive) differential band characteristic of the QAQB state given by an electrochromic shift on a C O group of the bacteriopheophytin HA located close to QA [37]. This makes it difficult to assess the real intensity of the 1726 cm1 band. Concerning the 1717 cm1 band, its presence in the first and second spectra recorded after the flash with the same amplitude suggest that already in the first spectrum Glu-L212 is completely protonated; however it cannot be excluded that transient contributions arising from other residues are present in the same region in the first spectrum, masking the real intensity of the Glu-L212 band. An interesting point concerns the absence in the first rapidscan spectrum of the positive band at 1689 cm1. Such band, as discussed before, was identified in wild-type RCs in D2O and possibly attributed to a transiently protonated side chain of a Glu or Asp (being the possible D2O correspondent of the transient 1707 cm1 band identified in H2O). Its absence would mean, if the above-mentioned interpretation is correct, that in the Asp-M17 ! Asn mutant such transient protonation does not take place or that at least it cannot be resolved by the rapidscan technique. It is interesting to note that in rapid-scan FTIR spectra of Asp-M17 ! Asn mutant RCs in H2O the 1707 cm1 band is also absent [21]. All these evidences suggest the possibility that the Asp-M17 residue could be responsible of the transient
A. Mezzetti, W. Leibl / Vibrational Spectroscopy 48 (2008) 126–134
1707 cm1 band. However, it should also be noted that in the first rapid-scan spectrum after the 1st flash on the AspL210 ! Asn/Asp-M17 ! Asn double mutant a positive band at 1705 cm1 is observed [21]. Therefore, the attribution of the 1707 cm1 band to the Asp-M17 should be considered, on the basis of all the available rapid-scan data, as a hypothesis. 3.4. Rapid-scan FTIR spectra on Cd2+-treated wild-type RCs All the data described above, together with several other experimental evidences, suggest that (1) the 1st electron transfer reaction between QA and QB is strongly coupled to proton transfer and that (2) this proton transfer reaction – at least in the wild-type RCs – proceed through a transient protonation of a Glu or Asp residue side chain. In this framework, it is interesting to investigate how a perturbation of the site of proton uptake (located in the Asp-H124, His-H126, His-H128 region) can influence both the QAQB ! QAQB electron transfer reaction and the proton pathway. As a perturbing agent, we have chosen Cd2+, which binds at the Asp-H124, His-H126, His-H128 residues and has a well-characterized inhibition effect of proton uptake [12,38]. The decrease in the rates of proton and electron transfer reactions in the RC upon Cd2+ binding has already been studied by UV–vis spectroscopy, proton uptake measurements [12] and photovoltage kinetics [38]. Compared to these methods, time-resolved FTIR provides not only the possibility of following the reaction through marker bands for QA, QB, QA, QB, GluL212 protonated side chain and protein rearrangement, but also to investigate the influence of Cd2+ on the transient protonation of a Glu or Asp residue characterized by the transient 1707 cm1 band described above. Results are shown in Fig. 7. The first scan (recorded between 4 ms and 29 ms after the laser flash) shows contributions arising from both the QA/QA state (positive band at 1467 cm1; negative band at 1670 cm1) and QB/QB state (positive band at 1479 cm1). This means that during the first scan we are monitoring the ongoing of the electron
transfer reaction. In the following spectrum, recorded between 77 ms and 102 ms, only bands characteristic of QBappear (e.g. band at 1479 (+) cm1). A positive band is also present at 1728 cm1, characteristic of Glu-L212 side chain protonation [17]. The following spectra do not show any appreciable further evolution. These data show that Cd2+ has a clear slowing effect on the electron transfer reaction. Unfortunately, also in this case it is very difficult to assess the presence and the intensity of the 1728 cm1 band in the first spectrum as it overlaps with other spectral features such as the 1735 ()/1727 (+) cm1 differential band characteristic of the QAQB state. The negative and quite broad band observed in the first scan at 1738 cm1 is probably given by such an effect (the shift compared to the 1735 cm1 band could possibly arise from a broadening effect due to the rapidscan technique combined with the overlapping rising 1728 cm1 positive band, although some spectral shift induced by the presence of Cd2+ cannot be excluded). Interestingly, there is no evidence for the 1707 cm1 band in any of the spectra.5 This is quite surprising, because we could expect that if Cd2+ binding slows down the electron transfer reaction, it should also slow down the proton transfer reaction, making the observation of the 1707 cm1 intermediate more easy (as in the case of the low-temperature experiments shown before). The absence of the 1707 cm1 band could be explained by the fact that Cd2+ binding affects the proton pathway, hampering the protonation of the Asp or Glu side chain responsible for the 1707 cm1. The crystal structure of the RC shows that Asp-M17 and Asp-L210 are extremely close to the binding site of Cd2+ [39], suggesting that the presence of the metal could modify their pKa and therefore their availability to take part in the proton transfer mechanism. Furthermore, recent calculations have shown that the effect of pKa modification induced by the presence of Cd2+ is strong and affects several other protonable amino acid side chains including Asp-L213, ˚ ) from the binding which is located relatively far away (12 A 2+ site of Cd [40]. These pKa shifts are probably responsible for a slow proton transfer to the QB site which in turn slows down also the electron transfer reaction [40]. It is important to mention that preliminary rapid-scan experiments on Ni2+-treated RCs (not shown) show a similar effect (i.e. absence of the 1707 cm1 band in all the spectra). Ni2+ binds to the RC at the His-H126 and AspM17 residues, very close to the Cd2+ binding site [40]. It is difficult to identify which is the amino acid whose side chain is responsible for the 1707 cm1 band, as all residues involved in the proton transfer pathway are possibly interconnected so that the presence of a metal or the mutation of an amino acid can in principle affect other residues. In the previous paragraph we have suggested that the 1707 cm1 band could arise from the Asp-M17 side chain. Nevertheless, more detailed investigations on mutant RCs will be necessary in order to unambiguously identify the residue responsible of the transient 1707 cm1 band.
5
2+
Fig. 7. Rapid-scan FTIR spectra recorded after the 1st flash in Cd -treated wild-type RCs. T = 281 K.
133
It should however be noted that, given the relatively high noise level, the presence of very small contributions of the 1707 cm1 band in the first spectrum cannot be totally excluded.
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4. Conclusions In this paper, we have shown that rapid-scan FTIR difference spectroscopy can be successfully used to investigate lightinduced reactions in bacterial RCs. From a methodological point of view, we have shown that quite simple strategies and/or experiment design can considerably increase the amount of information that can be obtained from spectral data. In particular, combination of experiments at different temperatures, in different media (H2O and D2O), on mutant RCs, or in presence of proton uptake inhibitors has allowed to extract from rapid-scan data new and interesting pieces of information concerning the proton and electron transfer reactions and the coupling between these two physicochemical processes. More precisely, we have shown that (1) the 1707 cm1 transient band observed after the 1st and the 2nd flashes is sensitive to temperature and to H/D exchange. This strongly suggests its identification with a Glu or Asp residue side chain transient protonation; (2) the QAQB ! QAQB electron transfer reaction in the AspM17 ! Asn mutant is slowed down by H/D exchange. This effect shows that in this mutant electron transfer is ratelimited by a process (involving proton movements in the protein) which is strongly influenced by H/D exchange; (3) consistent with previous UV–vis measurements, the QAQB ! QAQB electron transfer reaction is slowed down in presence of a proton uptake inhibitor (Cd2+); (4) in Cd2+-treated RCs the transient 1707 cm1 band is not observed, possibly because of a modification of the whole proton transfer pathway upon Cd2+ binding. Therefore this suggests its identification with a residue located on the proton transfer pathway. Acknowledgments The authors thank Prof. M.Y. Okamura and Prof. M.L. Paddock for the gift of the Asp-M17 ! Asn mutant RCs. A.M. thanks CEA/Saclay, the ‘‘Angelo Della Riccia’’ and ‘‘Guido Donegani’’ Foundations for partial financial support. References [1] P. Mitchell, J. Moyle, The role of ubiquinone and plastoquinone in chemiosmotic coupling between electron transfer and proton translocation, in: G. Lenaz (Ed.), Coenzyme Q, J Wiley & Sons, Chichester, 1985, p. 145. [2] J.P. Allen, G. Feher, T.O. Yeates, H. Komyia, D.C. Rees, Proc. Natl. Acad. Sci. U.S.A. 85 (1988) 8487. [3] U. Ermler, G. Fritzsch, S. Buchanan, H. Michel, Structure 2 (1994) 925. [4] G. Fritzsch, J. Koepke, R. Diem, A. Kuglstatter, L. Baciou, Acta Crystallogr. D: Biol. Crystallogr. 58 (2002) 1660. [5] M.Y. Okamura, M.L. Paddock, M.S. Graige, G. Feher, Biochim. Biophys. Acta: Bioenergetics 1458 (2000) 148. [6] M.L. Paddock, G. Feher, M.Y. Okamura, FEBS Lett. 555 (2003) 45. [7] M.S. Graige, G. Feher, M.Y. Okamura, Proc. Natl. Acad. Sci. U.S.A. 95 (1998) 11679. [8] M.H.B. Stowell, T.M. McPhillips, D.C. Rees, S.M. Soltis, E. Abresch, G. Feher, Science 276 (1997) 812.
[9] A. Remy, K. Gerwert, Nat. Struct. Biol. 10 (2003) 637. [10] M.S. Graige, M.L. Paddock, J.M. Bruce, G. Feher, M.Y. Okamura, J. Am. Chem. Soc. 118 (1996) 9005. [11] P.H. McPherson, M.Y. Okamura, G. Feher, Biochim. Biophys. Acta: Bioenergetics 1016 (1989) 289. [12] P. Adelroth, M.L. Paddock, L.B. Sagle, G. Feher, M.Y. Okamura, Proc. Natl. Acad. Sci. U.S.A. 97 (2000) 13086. [13] A. Barth, C. Zscherp, Quart. Rev. Biophys. 35 (2002) 369. [14] J. Breton, E. Naberdyk, Biochim. Biophys. Acta: Bioenergetics 1275 (1996) 84. [15] D.L. Thibodeau, E. Nabedryk, R. Hienerwadel, F. Lenz, W. Ma¨ntele, J. Breton, Biochim. Biophys. Acta: Bioenergetics 1020 (1990) 253. [16] E. Nabedryk, K.A. Bagley, D.L. Thibodeau, M. Bauscher, W. Ma¨ntele, J. Breton, FEBS Lett. 266 (1990) 59. [17] E. Nabedryk, J. Breton, R. Hienerwadel, C. Fogel, W. Ma¨ntele, M.L. Paddock, M.Y. Okamura, Biochemistry 34 (1995) 14722. [18] R. Hienerwadel, D.L. Thibodeau, F. Lenz, E. Nabedryk, J. Breton, W. Kreutz, W. Ma¨ntele, Biochemistry 31 (1992) 5799. [19] R. Hienerwadel, S. Grzybek, C. Fogel, W. Kreutz, M.Y. Okamura, M.L. Paddock, J. Breton, E. Nabedryk, W. Ma¨ntele, Biochemistry 34 (1995) 2832. [20] J.-R. Burie, W. Leibl, E. Nabedryk, J. Breton, Appl. Spectrosc. 47 (1993) 1401. [21] A. Mezzetti, E. Nabedryk, J. Breton, M.Y. Okamura, M.L. Paddock, G. Giacometti, W. Leibl, Biochim. Biophys. Acta: Bioenergetics 1553 (2002) 320. [22] A. Mezzetti, W. Leibl, J. Breton, E. Nabedryk, FEBS Lett. 537 (2003) 161. [23] A. Mezzetti, W. Leibl, Eur. Biophys J. 34 (2005) 921. [24] R. Brudler, K. Gerwert, Photosynth. Res. 55 (1998) 261. [25] S. Hermes, J.M. Stachnik, D. Onidas, A. Remy, E. Hofma, K. Gerwert, Biochemistry 45 (2006) 13741. [26] L. Blanchet, A. Mezzetti, C. Ruckebusch, J.P. Huvenne, A. de Juan, Anal. Bioanal. Chem. 387 (2007) 1863. [27] M.L. Paddock, G. Feher, M.Y. Okamura, Proc. Natl. Acad. Sci. U.S.A. 97 (2000) 1548. [28] R.A. Isaacson, F. Lendzian, E.C. Abresch, W. Lubitz, G. Feher, Biophys. J. 69 (1995) 311. [29] J. Breton, C. Berthomieu, D.L. Thibodeau, E. Nabedryk, FEBS Lett. 278 (1991) 257. [30] J. Breton, D.L. Thibodeau, C. Berthomieu, W. Ma¨ntele, A. Verme´glio, E. Nabedryk, FEBS Lett. 288 (1991) 109. [31] E. Nabedryk, Light-induced Fourier transform infrared difference spectroscopy of the primary donor in photosynthetic reaction centers, in: H. Mantsch, D. Chapman (Eds.), Infrared Spectroscopy of Biomolecules, Wiley–Liss Inc., 1996, p. 39. [32] J. Breton, J.-R. Burie, C. Berthomieu, D.L. Thibodeau, S. Andrianambinintsoa, D. Dejonghe, G. Berger, E. Nabedryk, Light-induced charge separation in bacterial photosynthetic reaction centers monitored by FTIR difference spectroscopy: the QA vibrations, in: J. Breton, A. Verme´glio (Eds.), The Photosynthetic Bacterial Reaction Center, vol. II, Plenum Press, New York, 1992, p. 155. [33] A. Barth, Prog. Biophys. Mol. Biol. 74 (2000) 141. [34] M.L. Paddock, P. Aderlroth, C. Chang, E.C. Abresch, G. Feher, M.Y. Okamura, Biochemistry 40 (2001) 6893. [35] L.I. Krishtalik, Biochim. Biophys. Acta: Bioenergetics 1458 (2000) 6. [36] E. Nabedryk, J. Breton, M.Y. Okamura, M.L. Paddock, Biochemistry 40 (2001) 13826. [37] J. Breton, E. Nabedryk, J.P. Allen, J.C. Williams, Biochemistry 36 (1997) 4515. [38] S. Keller, J.T. Beatty, M.L. Paddock, J. Breton, W. Leibl, Biochemistry 40 (2001) 429; S. Keller, J.T. Beatty, M.L. Paddock, J. Breton, W. Leibl, Biochemistry 387 (2007) 1863. [39] H.L. Axelrod, E.C. Abresch, M.L. Paddock, M.Y. Okamura, G. Feher, Proc. Natl. Acad. Sci. U.S.A. 97 (2000) 1542. [40] H. Isikita, E.-W. Knapp, Proc. Natl. Acad. Sci. U.S.A. 102 (2005) 16215.
Vibrational Spectroscopy 48 (2008) 126–134 www.elsevier.com/locate/vibspec
Proton and electron transfer in wild-type and mutant reaction centers from Rhodobacter sphaeroides followed by rapid-scan FTIR spectroscopy Alberto Mezzetti a,b,*, Winfried Leibl b,c a
Laboratoire de Spectrochimie Infrarouge et Raman UMR CNRS 8516, Universite´ de Sciences et Technologies de Lille, Bat C5, Cite´ Scientifique, 59655 Villeneuve d’Ascq, France b CEA, iBiTecS, Service de Bioe´nenerge´tique Biologie Structurale et Me´canismes (SB2SM), F-91191 Gif-sur-Yvette, France c CNRS, URA 2096, F-91191 Gif-sur-Yvette, France Received 17 July 2007; received in revised form 20 January 2008; accepted 29 January 2008 Available online 15 February 2008
Abstract Rapid-scan FTIR difference spectroscopy was used to investigate proton and electron transfer reactions in photosynthetic reaction centers from Rhodobacter sphaeroides. Experiments at different temperatures and in the presence of D2O have provided strong indication that a transient band at 1707 cm1 previously identified after both the 1st and the 2nd flashes [A. Mezzetti, W. Leibl, Eur. Biophys. J. 34 (2005) 921] is given by a transient protonation of the side chain of a Asp or Glu residue situated on the proton transfer pathway from the cytoplasm to the QB site. Experiments in D2O on a Asp-M17 ! Asn mutant reaction center, where the proton and electron transfer reactions are slowed down compared to the wild-type, showed that the kinetic isotope effect induced by H/D exchange slows down the electron transfer reaction after the 1st flash, confirming the strong coupling between proton and electron transfer. Rapid-scan FTIR experiments on Cd2+-treated reaction centers, in agreement with previous UV–vis measurements [P. Adelroth, M.L. Paddock, L.B. Sagle, G. Feher, M.Y. Okamura, Proc. Natl. Acad. Sci. U.S.A. 97 (2000) 13086], showed that upon addition of Cd2+, which inhibits proton uptake, the QAQB ! QAQB reaction is slowed. Interestingly, the transient 1707 cm1 band is not visible in the first spectrum recorded early after the flash. This strongly suggests its identification with a residue situated on the proton transfer pathway, which is perturbed upon metal cation binding. # 2008 Elsevier B.V. All rights reserved. Keywords: Rapid-scan FTIR; Rhodobacter sphaeroides; Photosynthetic reaction center; FTIR difference spectroscopy; Proton transfer; Electron transfer
1. Introduction Photosynthetic reaction centers (RCs) from purple bacteria are membrane enzymes which act as light-driven quinone reductases. Absorption of a photon entails an ultrafast charge separation between the so-called primary electron donor P (a special pair of bacteriochlorophyll-a molecules) and the primary acceptor QA, a quinone molecule strongly bound to the protein. A second quinone molecule, called QB, accepts two
* Corresponding author at: Laboratoire de Spectrochimie Infrarouge et Raman UMR CNRS 8516, Universite´ de Sciences et Technologies de Lille, Bat C5, Cite´ Scientifique, 59655 Villeneuve d’Ascq, France. Fax: +33 3 20 43 66 03. E-mail address: [email protected] (A. Mezzetti). 0924-2031/$ – see front matter # 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.vibspec.2008.01.019
electrons from QA in two consecutive photochemical events, as well as two protons from the cytoplasm. Protons are transferred to the QB site through a pathway formed by amino acid side chains and water molecules. The formed ubiquinol molecule QBH2 leaves the RC and is replaced by an oxidised quinone coming from the quinone pool present in the membrane. The ubiquinol is then reoxidised by another membrane enzyme, called the cytochrome bc1 complex. The overall effect of this reaction cycle is to move protons from the cytoplasm to the periplasm. The formed proton gradient is then used to synthesise ATP, necessary to power the metabolic reactions in the cell. Proton and electron transfer reactions, especially those involving quinone molecules, play a key role in bioenergetics [1]. Compared to other enzymes, in bacterial photosynthetic RCs the reactions can be easily triggered by short laser pulses.
A. Mezzetti, W. Leibl / Vibrational Spectroscopy 48 (2008) 126–134
This makes photosynthetic RCs ideal systems to investigate general principles of quinone redox chemistry as well as proton and electron transfer in bioenergetic systems. The structure of the RC from Rhodobacter (Rb.) sphaeroides ˚ has been determined by X-ray crystallography with up to 2 A resolution [2–4] leading to a detailed knowledge of the positions of pigments, cofactors and amino acids, as well as on their relative distance and orientation. This makes possible to give a much deeper interpretation of data coming from spectroscopic techniques, in order to fully understand the mechanisms of the reactions taking place in the RCs. In addition, for Rb. sphaeroides site-directed mutagenesis is wellestablished. Knowledge of the structure also makes possible to change specific amino acids in key positions of the RC. Mutant RCs have indeed provided most of the recent relevant data on the working mechanism of the RC, especially concerning the proton transfer pathways (see [5,6] for recent reviews). It is now generally accepted that in isolated RCs the first electron transfer from QA to QB does not involve protonation of QB, which occurs only on the second reduction step. However, the electron transfer step is accompanied by a substoichiometric proton uptake from the cytoplasm towards the interior of the protein. In addition, driving force assay methods have demonstrated that the QAQB ! QAQB reaction is not rate-limited by an electron transfer but by another step [7]. Even though different processes have been proposed as the ratelimiting step [5,8], the implication of a proton transfer is strongly suggested by recent studies [6,9]. The second electron transfer between the two quinones leads to the formation of a QBH state in less than 1 ms. This electron transfer is coupled to, but not limited by, proton uptake from the cytoplasm [10]. Subsequently, QH is rapidly protonated and the formed QBH2 leaves the RC [11]. There is no definite agreement on the detailed mechanism of the two electron transfer reactions. Nevertheless, a dominant proton pathway from the cytoplasm to the QB site has been identified [5,6] which is active for proton transfer towards the QB site upon both electron transfer reactions. In both cases, protons are taken up by the His-H126 and His-H128 residues (Fig. 1), which in fact are part of the binding site of the proton transfer inhibitors Zn2+ and Cd2+ (Ni2+, another proton transfer inhibitors, binds at His-H126 and Asp-M17) [12]. Located between the entry point and the QB site are Asp-L210 and AspM17. These two residues play a crucial role in the transfer of both protons. Also the Asp-L213 residue is believed to be part of the proton transfer pathway. At or near Asp-L213, the pathways for the two protons diverge (see Fig. 1). Proton transfer to Glu-L212 (associated with the first electron transfer) occurs through intervening water molecules. H+ transfer directly to QB (associated to the second electron transfer) occurs through the Ser-L223 side chain. Water molecules located between the amino acid side chains are likely to act as connectors. FTIR difference spectroscopy has been largely used to investigate structure-function relationships in Rb. sphaeroides RCs and chromatophores, relying on the capability of the technique to monitor simultaneously reaction-induced changes
127
Fig. 1. Superimposed on part of the crystal structure are the proton transfer pathways in Rhodobacter sphaeroides RCs. See text for further details. Figure is adapted from [36].
in both the cofactors and the proteins [13]. A large amount of data has been obtained by static FTIR difference spectroscopy leading to the precise identification of IR marker bands for specific redox states of cofactors ([14] and references therein), protein rearrangements [15,16] and protonation of amino acid side chains [17]. The availability of these marker bands has prompted a series of time-resolved IR difference spectroscopy investigations. Whereas early investigations used mainly single wavelength techniques [18,19], in more recent years timeresolved FTIR techniques have been used. After the pioneering experiments of Thibodeau et al. [15] and Burie et al. [20], Rb. sphaeroides RCs and chromatophores have been studied by rapid-scan FTIR [21–23] and step-scan FTIR [9,24]. Recently, an investigation combining both the rapid-scan and the stepscan technique has been reported [25]. It is also worth mentioning that very recently a chemometric approach to analyse time-resolved FTIR difference spectra of Rb. sphaeroides chromatophores has been proposed [26]. In this paper, we report new rapid-scan FTIR data aimed to better understand the proton and electron transfer reactions taking place in isolated Rb. sphaeroides RCs. The simultaneous investigation of the reactions in different conditions (different temperatures, different media—H2O and D2O, use of native and site-directed mutant RCs, presence of proton uptake inhibitors) provided new insight into the coupling between electron and proton transfer reactions. 2. Experimental 2.1. Sample preparation The site-directed mutation Asp-M17 ! Asn was constructed as described previously [27]. RCs from Rb. sphaeroides R26 and the mutant were isolated in 15 mM Tris–HCl pH 8, 0.025% lauryl dimethylamide-Noxide (LDAO), as described in [28]. The samples for FTIR measurements were prepared as previously described [29,30].
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Briefly, RCs were deposited on a CaF2 window and dried under argon. Before complete dryness, the RC film was re-hydrated with Tris buffer (see below). To guarantee full occupancy of the QB site, a fivefold excess of UQ6 (ubiquinone-6, Sigma) was added to the RC suspension. To ensure fast reduction of P+ external donors were added. Two different mixtures were used, following published protocols [23]: (i) ferrocyanide (250 mM) and tetramethyl-p-phenylenediamine (TMPD; 50 mM) buffered with Tris (pH 7; 100 mM) and (ii) 10 mM Na-ascorbate and 20 mM 2,3,5,6-tetramethyl-p-phenylenediamine (DAD) buffered at pH 8. The choice of the redox mediators determines the re-oxidation time of QBH2 between averaging cycles. A second CaF2 window was used to squeeze the sample, in order to yield an absorbance in the amide I region of the spectrum of 0.6–0.9 a.u. 2.2. FTIR difference spectroscopy experiments Rapid-scan measurements were performed using the experimental parameters as described previously [23]. A Bruker IFS 88 FTIR spectrometer equipped with a photoconductive MCT-A detector and with Opus software was used. Measurements were performed at 285 1 K, 281 1 K, 268 1 K and 260 1 K using a temperature-controlled N2 cryostat. The photoreactions were triggered by a saturating flash from a frequency-doubled Nd:YAG laser (7 ns, 20 mJ, Quantel, France). The experimental scheme used varied slightly according to the particular sample studied. For wild-type, untreated RCs, 40 interferograms were recorded in the dark and averaged. Then a first laser flash was fired and an interferogram was recorded between 2 ms and 27 ms to monitor the time evolution of the system after the flash. 73 ms after the 1st flash, a 2nd flash was fired (see
Fig. 2B). An interferogram was then recorded between 2 ms and 27 ms after the 2nd flash. Then four interferograms were recorded at increasing delay to monitor the process of ubiquinol formation. Subsequent interferograms were averaged in groups of five. The results from 3200 cycles (obtained on two different samples) were averaged to improve the signal-to-noise ratio. For mutant and Cd2+-treated wild-type RCs, 40 interferograms were recorded in the dark and averaged. Then a first laser flash was fired and an interferogram was recorded between 4 ms and 29 ms to monitor the time evolution of the system after the flash. Then four interferograms were recorded at increasing delay to monitor the process of ubiquinol formation (see Fig. 2A). Subsequent interferograms were averaged in groups of five. The results from 2000 cycles (obtained on two different samples) were averaged to improve the signal-to-noise ratio. Between cycles an appropriate delay time (3 min for the samples with DAD and Na-ascorbate as redox mediators; a time ranging from 30 s to 3 min, according to the temperature, for the samples with ferrocyanide and TMPD as redox mediators) allowed a complete relaxation of the system due to re-oxidation of QH2 by the external redox chemicals. The necessary relaxation time was determined by investigating the decay of the differential spectra at long times. At the end of the experiments, the stored interferograms were Fourier transformed to give the corresponding single beam spectra using the Opus software, applying the Mertz phase correction and the Blackman-Harris 3-term apodization function. Difference spectra at various times after the laser flash were calculated according to the formula DA(t) = log[S(t)/S(0)], where S(0) is the single beam spectrum obtained from the averaged interferogram recorded before the 1st flash and S(t) is the single beam spectrum at time t after the 1st or the 2nd flashes.
Fig. 2. Scheme for (A) experiments with 1 triggering flash; (B) experiments with 2 triggering flashes.
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3. Results and discussion 3.1. Rapid-scan FTIR spectra after the 1st and 2nd flashes at different temperatures Time-resolved FTIR difference spectra after one and two consecutive flashes recorded at 285 K, 268 K and 260 K are shown in Figs. 3 and 4. In these experiments ferrocyanide and TMPD were used as external donors in order to quickly reduce P+, which would give rise to IR absorption bands which are much more intense (by a factor of 5) than quinone, semiquinone and protein bands [31]. The ferrocyanide/TMPD mixture can reduce P+ more quickly than the conventional DAD/ascorbate mix used in most steady-state measurements and previously used also in rapid-scan FTIR measurements [21–23,25,26]. Moreover, ferrocyanide/TMPD ensures fast P+ reduction also at low temperature (260 K and 268 K). Although TMPD redox changes are known to give relatively intense IR signals at 1620 cm1 (negative signal), 1546 cm1 (positive signal), 1520 cm1 (negative), 1431 cm1 (positive) and 1382 cm1 (positive) [32] (which in several cases can be a serious drawback), no signals arising from TMPD+/TMPD are expected in the 1650–1750 cm1 region. Therefore, peaks in this spectral region can be safely attributed to protein and/or cofactor contributions. In Fig. 3 are reported the spectra recorded in the first scan after the 1st flash (between 2 ms and 27 ms after the flash) at the three temperatures, corresponding to QB/QB difference spectra. In fact bands characteristic of QB (positive band at 1479 cm1) of disappearing QB (negative band at 1264 cm1) and of protonation of the Glu-L212 side chain (1728 cm1) are visible at all three temperatures. Bands reflecting at least partial contributions from TMPD+/TMPD signals are visible at 1620 cm1 (negative), 1541 cm1 (positive) and 1523 cm1 (negative). These TMPD+/TMPD bands overlap with contributions from the RC; this is possibly the reason why they appear at
Fig. 3. Rapid-scan FTIR difference spectra at different temperatures recorded between 2 ms and 27 ms after the 1st flash in presence of ferrocyanide and TMPD in wild-type RCs. Bands arising – at least partially – from TMPD+/ TMPD contributions are marked by an asterisk.
Fig. 4. Rapid-scan FTIR difference spectra at different temperatures recorded between 2 ms and 27 ms after the 2nd flash in presence of ferrocyanide and TMPD in wild-type RCs. Bands arising – at least partially – from TMPD+/ TMPD contributions are marked by an asterisk.
a slightly different wavenumbers than found in electrochemically generated TMPD+/TMPD FTIR difference spectra of pure TMPD solutions [32]. Smaller TMPD+/TMPD contributions are present in our spectra but do not emerge as clear positive or negative bands as they are hidden by stronger spectral contributions from the protein and the cofactors. In addition, a positive band at 1707 cm1 is present at 260 K and 268 K, but absent at 285 K. This band has been previously observed at 281 K (using DAD and ascorbate as redox mediators) and 268 K (using ferrocyanide and TMPD) [23]. Experiments where more scans were measured after the 1st flash (data not shown; see also [23] where this experiment was carried out at 281 K) showed that this band quickly decays in intensity (it is absent in the following scans), in contrast to the QB band (1479 cm1) and the protonated Glu-L212 side chain band (1728 cm1) which are more stable. In our previous paper, this was interpreted as an indication that the 1707 cm1 band corresponds to a transient event associated with the formation of QB [23]. In Fig. 4 are shown the spectra recorded in the first scan (between 2 ms and 27 ms) after the 2nd flash at 285 K, 268 K and 260 K. They correspond to a QB(double reduced)/QB state. They all show a positive band at 1479 cm1, but with decreased intensity and increased broadness compared to the spectrum recorded immediately after the 1st flash. This band was previously reported at 281 K and attributed to undesired formation of QB in a portion of RCs [23]. Unfortunately the marker bands for QH2 and/or QBH formation overlap with the strong positive bands from TMPD+. Indications for QH2 and/or QBH formation can however be seen at 1490 cm1 (shoulder) as well as from the strong decrease in intensity of the 1479 cm1 band after the 2nd flash. Also the band at 1728 cm1 is present (but with decreased amplitude compared to the spectra recorded immediately after the 1st flash). The band at 1707 cm1 is present in the three spectra but with different intensities: it is strong at 268 K and 260 K, but weak at
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285 K. Experiments where more scans were measured after the 2nd flash (data not shown; see also [23] where this experiment was carried out at 281 K and 268 K) indicate that this band quickly decays in intensity. By comparing the spectra at the three temperatures after both the 1st and the 2nd flashes it clearly emerges that the intensity of the positive band at 1707 cm1 decreases as the temperature increases, whereas almost all other peaks are present with roughly the same intensity at all temperatures. If we focus our attention on the 285 K rapid-scan FTIR spectra, we notice that the 1707 cm1 band is absent in the first scan after the 1st flash, whereas it is present but very weak in the first scan recorded after the 2nd flash. As reported above, this band has previously been observed at 268 K and 281 K after both the 1st and the 2nd flashes and tentatively attributed to a transient protonation of a Glu or Asp side chain [23]. The absence (or very small amplitude) of this transient band at 285 K can indeed be easily explained by an effect of temperature on the reaction kinetics. It has been reported that proton transfer reactions in Rb. sphaeroides RC are slowed down as the temperature is lowered [9,21]. Therefore the decrease in intensity of the 1707 cm1 band with increasing temperature is compatible with (and supportive of) its assignment to a transiently protonated side chain of a Glu or Asp residue. In fact, it can be argued that such a transient protonation is visible in the first rapid-scan FTIR spectrum at 260 K and 268 K but becomes very weak (after the 2nd flash) or no longer visible (after the 1st flash) in the same time window (2–27 ms after the flash) at 285 K, because its decay is faster than the time resolution of the technique. We recall that rapidscan experiments at 281 K [23] (but performed using DAD/ ascorbate as redox mediators and at pH 8 instead of 7) showed a weak – but detectable – transient band at 1707 cm1 after both the 1st and the 2nd flashes. Although direct comparison with the present results is questionable due to different experimental conditions (notably the used redox mediators and the pH), it appears that the intensity of the 1707 cm1 band lies between those obtained at 285 K and those obtained at 268 K (not shown) which is consistent with the temperature influence deduced above on the 1707 cm1 band deduced above. A transient 1707 1 cm1 band has also been observed at 278 K by single wavelength time-resolved IR spectroscopy [19] and by time-resolved step-scan FTIR spectroscopy [9].1
3.2. Rapid-scan FTIR spectra on wild-type RCs in D2O: H/D exchange effects on band position In order to confirm the attribution of the 1707 cm1 band to a transiently protonated Glu or Asp side chain, we performed experiments in D2O, as in these conditions we should observe a
Fig. 5. Rapid-scan FTIR spectra recorded on wild-type RCs. From top to bottom: spectra recorded between 2 ms and 27 ms after the 1st flash in D2O at 285 K (continuous line) and H2O at 260 K (dotted line); spectra recorded between 2 ms and 27 ms after the 2nd flash in D2O at 285 K (continuous line) and H2O at 260 K (dotted line).
downshift of the band due to H/D exchange [33]. Rapid-scan FTIR difference spectra recorded in D2O after the 1st and the 2nd flashes at 285 K2 are shown in Fig. 5. Ferrocyanide and TMPD were used as external donors, as in the case of rapidscan FTIR spectra recorded in H2O (reported in Figs. 3 and 4). The first spectrum recorded after the 1st flash (Fig. 5) is indicative of QB reduction (positive band at 1479 cm1; not shown) as well as protonation of the Glu-L212 side chain (band at 1728 cm1); on the other hand the 1707 cm1 band is very weak. Interestingly, we observe a new positive band at 1717 cm1. A positive band at 1717 cm1 has been previously reported in steady-state QB/QB FTIR difference spectra of wild-type RCs and interpreted as consequence of H/D exchange
2
1
It should be noticed that, in contrast with [9,19] in the present experiments the proton transfer reaction was investigated after fast and complete reduction of the oxidised primary donor P. This eliminates disturbing spectral contributions arising from the intense P+/P bands and better reproduces the situation occurring under physiological conditions.
As explained in the following, recording of D2O spectra at 285 K is suggested by kinetic considerations. However, the rate of P+ reduction by external donors seems to be severely reduced in D2O at low temperatures. We observed that below 285 K complete P+ reduction before the beginning of the first scan after both the 1st and the 2nd flashes is no longer efficient. To obtain fast and highly reproducible reduction of P+ the temperature had to be set to 285 K.
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at the protonated side chain of Glu-L212 [17]. After the 2nd flash, a small contribution is present at 1728 cm1 whereas the band at 1707 cm1 is again very weak. It is useful to compare these data with those obtained in H2O in order to point out possible H/D exchange-induced band shifts. However, comparison with H2O spectra recorded at the same temperature is not a proper choice, because we have to take into account a probable kinetic isotope effect. Indeed, the proton transfer reaction associated to the QAQB ! QAQB electron transfer shows a kinetic isotope effect of 4.9 in wildtype Rb. sphaeroides RCs [25]. Therefore we expect that rapidscan spectra recorded at 285 K in H2O and D2O in the first scan after the 1st flash do not represent the very same situation. In other words, if we consider the 2–27 ms time window to roughly correspond to a snapshot at 14.5 ms after the flash, in the two cases different stages of the reaction are monitored, as the proton transfer is expected to proceed at a significantly higher rate in H2O compared to D2O. On the other hand, if we now consider the data obtained in H2O presented above, we have strong indications – consistent with previous observations [9,21] – for an increase of the rate of the proton transfer with the temperature. Indeed, a 20-degree difference in temperature (298 K vs. 278 K) was reported to correspond to a 3.5-fold difference in kinetics of the proton transfer reaction coupled to the QA QB ! QAQB electron transfer reaction [9]. This is consistent with the observed behaviour found in the Asp-L210 ! Asn/Asp-M17 ! Asn double mutant (compare Mezzetti et al. [21] and Paddock et al. [34]) where a 13-degree difference in temperature (294 K vs. 281 K) was found to correspond to a 4-fold difference in kinetics of the QAQB ! QAQB electron transfer reaction and of the associated proton transfer. It can therefore be argued that for the proton transfer reaction coupled to the first electron transfer between QA and QB it is more appropriate to compare H2O data at 260 K with D2O data at 285 K, as the temperature-induced deceleration of proton transfer in H2O roughly compensates the kinetic isotope effect in D2O. For the proton transfer reaction coupled to the 2nd electron transfer between QA and QB no literature data are available for a possible kinetic isotope effect, even though such an effect is likely to occur, as it normally affects proton-transfer reactions in protein [35]. On the other hand, the rapid-scan data shown above clearly indicate that the process involving the formation of the 1707 cm1 is slowed down as the temperature is lowered. Therefore the comparison between the 285 K spectrum in D2O and the 260 K rapid-scan spectrum in H2O seems proper also when spectra recorded in the first scan after the 2nd flash are compared. In Fig. 5 (dotted lines) we have reported the rapid-scan spectra after the 1st and the 2nd flashes in H2O at 260 K to allow a direct visual comparison.3
3 Intensities have been normalized to yield identical amplitudes of the 1264 cm1 band which is given by a C–O–CH3 mode of a methoxy group.
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Even though a more accurate approach would require calculation of double difference spectra (D2O minus H2O),4 some features are clearly visible from the direct comparison of the D2O and H2O spectra. The 1707 cm1 band is clearly sensitive to H/D exchange, as its intensity is considerably lower in D2O. This strengthens the identification of this band as the C O stretching of a protonable residue (Asp or Glu). One possibility is that in D2O it downshifts at 1689 cm1 (compare the spectra in H2O and D2O after both the 1st and the 2nd flashes). However, the 1689 cm1 band could also be (at least partially) given by the downshifting of the 1695 cm1 band. Another interesting feature is the intense negative band at 1687 cm1 present in spectra in H2O recorded immediately after both the 1st and the 2nd flashes. Whereas this band is absent in D2O spectra, in H2O it was found to be still present also at long delays after the 1st [21] and the 2nd flashes [23]. It has been tentatively assigned to an amide I contribution [36]. 3.3. Rapid-scan FTIR spectra on wild-type and mutant RCs in D2O: kinetic isotope effect on proton and electron transfer Comparison of time-resolved FTIR difference spectra in H2O and D2O can be used also to investigate the possible consequences of an isotope kinetic effect. In fact, IR spectroscopy can allow to investigate other possible slowing effects due to H/D exchange. We recall that in Rb. sphaeroides RCs IR marker bands are available for QA, QB, QA, QB [14], QH2 [22,23], protein rearrangements [15,16], and protonated side chain of amino acids [17]. Compared to normal UV–vis monitoring of the QA QB ! QAQB reaction, where global chromophore changes are observed, IR has the definite advantage of providing specific bands for almost any of the cofactors and molecular groups taking part in the reaction; furthermore, also the effect of H/D exchange on the position of these bands has been investigated [17]. Of course for the identification of a kinetic isotope effect we want to compare the kinetics for marker bands in H2O and D2O (possible shifts in marker band position can be easily taken into account relying on the existing literature). For this reason, the spectra to be compared must be recorded at the same temperature. As several evidences suggest that the first electron transfer between QA and QB is strongly coupled to a proton transfer reaction [5], it is possible that the observed decreased rate of 4 Calculations of double difference spectra (D2O minus H2O) is a common procedure in FTIR difference spectroscopy that can provide precise indication on H/D exchange-induced shift of bands. However this procedure requires (1) that the spectra are recorded under exactly the same conditions (except for the nature of the surrounding medium) and (2) that the spectra show an excellent signal-to-noise ratio. In time-resolved FTIR these two requirements are difficult to meet because of the intrinsic limitation in signal-to-noise ratio and because it is difficult to take properly into account the exact influence of the kinetic isotope effect (and therefore to choose the right pair of spectra for the subtraction procedure).
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proton transfer in D2O entails also a decreased rate of the electron transfer reaction. We recall that for rapid-scan spectra recorded in H2O no indication of an ongoing electron transfer is visible, even at low temperatures: in other words, the electron transfer reaction could not be resolved. For this reason we looked in the rapid-scan spectra in D2O at the spectral region where marker bands for the QAQB state lies, notably the intense positive band at 1467 cm1 [14] and the negative band at 1670 cm1 [15]. Unfortunately no traces of QAQB state can be found on the first rapid-scan spectrum recorded between 2 ms and 27 ms after the 1st flash (data not shown). Therefore, apparently in wild-type RCs at 285 K the kinetic isotope effect is incapable to slow down the electron transfer reaction between the two quinones sufficiently enough to be resolved by the rapid-scan technique. On the other hand, it is not possible to lower the temperature as in this case fast reduction of P+ is no longer efficient. We therefore looked for a mutant RC where the proton transfer coupled to the QAQB ! QAQB electron transfer reaction is already slowed down in H2O, but not enough to let QA contributions appear in the first scan. We focused our attention on an Asp-M17 ! Asn mutant which has been already investigated by UV–vis and proton uptake measurements [34], as well as static and rapid-scan FTIR [36,21]. This system is indeed ideal as in rapid-scan FTIR spectra on H2O no traces of QA were detectable in the first spectrum after the laser flash [21]. We performed some measurements on this mutant in D2O at 281 K and pH 8 (see spectra in Fig. 6), i.e. the very same conditions of temperature and pH of the published data of the Asp-M17 ! Asn mutant in H2O [21]. Also the timing of the experiment was set to be identical to the one for H2O [21], i.e. the first scan was recorded between 4 ms and 29 ms after the flash. Very interestingly, intense marker bands for the QAQB state are visible at 1467 cm1 and 1670 cm1 in the first scan after the 1st flash (Fig. 6, upper trace). Also the marker band for QB is present in the first scan (see band at 1479 cm1). Therefore, this spectrum reflects a situation in
Fig. 6. (A) Rapid-scan FTIR spectra on the Asp-M17 ! Asn mutant in D2O, T = 281 K. (B) Rapid-scan FTIR spectra on the Asp-M17 ! Asn mutant in H2O, T = 281 K; adapted from [21].
time where the electron transfer reaction is about half-way advanced. This means that the presence of D2O, combined with the 8-fold decrease caused by the mutation of the M17 amino acid [34], is capable to slow down the reaction enough to permit to observe intense QAQB state marker bands in the first scan after the laser flash. Such spectral contributions are absent in the following scans, where only bands characteristic of QB are present (see Fig. 6). Therefore, in the Asp-M17 ! Asn mutant D2O clearly affects the QAQB ! QAQB electron transfer reaction. The intrinsic mechanism of mono-electronic oxidation of QA and reduction of QB does not imply any protonation or deprotonation of the semiquinone or quinone. This means that the electron transfer reaction is rate-limited by an ‘‘external’’ step which is sensitive to H/D exchange. An interpretation would be to assume that the electron transfer is rate-limited by a proton transfer step. However, the issue on the rate-limiting step for the QAQB ! QAQB reaction is still under debate, and other ‘‘gating’’ steps sensitive to an isotopic effect are possibly involved. Nevertheless, the present results suggest a strong coupling between the QAQB ! QAQB reaction and proton transfer reactions. Unfortunately, no clear conclusions can be drawn concerning the protonation state of the Glu-L212 side chain in the first spectrum. In the following spectra, this residue is responsible for the positive bands at 1726 cm1 and 1717 cm1, as determined by steady-state FTIR difference measurements [36]; these two bands arise from the partial deuteration of the Glu-L212 residue [17,36], even though in steady-state spectra their position was slightly different (1728 cm1 and 1716 cm1). In the first spectrum (recorded between 4 ms and 29 ms after the flash), the 1726 cm1 band is overlapping with a 1735 cm1 (negative)/1727 cm1 (positive) differential band characteristic of the QAQB state given by an electrochromic shift on a C O group of the bacteriopheophytin HA located close to QA [37]. This makes it difficult to assess the real intensity of the 1726 cm1 band. Concerning the 1717 cm1 band, its presence in the first and second spectra recorded after the flash with the same amplitude suggest that already in the first spectrum Glu-L212 is completely protonated; however it cannot be excluded that transient contributions arising from other residues are present in the same region in the first spectrum, masking the real intensity of the Glu-L212 band. An interesting point concerns the absence in the first rapidscan spectrum of the positive band at 1689 cm1. Such band, as discussed before, was identified in wild-type RCs in D2O and possibly attributed to a transiently protonated side chain of a Glu or Asp (being the possible D2O correspondent of the transient 1707 cm1 band identified in H2O). Its absence would mean, if the above-mentioned interpretation is correct, that in the Asp-M17 ! Asn mutant such transient protonation does not take place or that at least it cannot be resolved by the rapidscan technique. It is interesting to note that in rapid-scan FTIR spectra of Asp-M17 ! Asn mutant RCs in H2O the 1707 cm1 band is also absent [21]. All these evidences suggest the possibility that the Asp-M17 residue could be responsible of the transient
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1707 cm1 band. However, it should also be noted that in the first rapid-scan spectrum after the 1st flash on the AspL210 ! Asn/Asp-M17 ! Asn double mutant a positive band at 1705 cm1 is observed [21]. Therefore, the attribution of the 1707 cm1 band to the Asp-M17 should be considered, on the basis of all the available rapid-scan data, as a hypothesis. 3.4. Rapid-scan FTIR spectra on Cd2+-treated wild-type RCs All the data described above, together with several other experimental evidences, suggest that (1) the 1st electron transfer reaction between QA and QB is strongly coupled to proton transfer and that (2) this proton transfer reaction – at least in the wild-type RCs – proceed through a transient protonation of a Glu or Asp residue side chain. In this framework, it is interesting to investigate how a perturbation of the site of proton uptake (located in the Asp-H124, His-H126, His-H128 region) can influence both the QAQB ! QAQB electron transfer reaction and the proton pathway. As a perturbing agent, we have chosen Cd2+, which binds at the Asp-H124, His-H126, His-H128 residues and has a well-characterized inhibition effect of proton uptake [12,38]. The decrease in the rates of proton and electron transfer reactions in the RC upon Cd2+ binding has already been studied by UV–vis spectroscopy, proton uptake measurements [12] and photovoltage kinetics [38]. Compared to these methods, time-resolved FTIR provides not only the possibility of following the reaction through marker bands for QA, QB, QA, QB, GluL212 protonated side chain and protein rearrangement, but also to investigate the influence of Cd2+ on the transient protonation of a Glu or Asp residue characterized by the transient 1707 cm1 band described above. Results are shown in Fig. 7. The first scan (recorded between 4 ms and 29 ms after the laser flash) shows contributions arising from both the QA/QA state (positive band at 1467 cm1; negative band at 1670 cm1) and QB/QB state (positive band at 1479 cm1). This means that during the first scan we are monitoring the ongoing of the electron
transfer reaction. In the following spectrum, recorded between 77 ms and 102 ms, only bands characteristic of QBappear (e.g. band at 1479 (+) cm1). A positive band is also present at 1728 cm1, characteristic of Glu-L212 side chain protonation [17]. The following spectra do not show any appreciable further evolution. These data show that Cd2+ has a clear slowing effect on the electron transfer reaction. Unfortunately, also in this case it is very difficult to assess the presence and the intensity of the 1728 cm1 band in the first spectrum as it overlaps with other spectral features such as the 1735 ()/1727 (+) cm1 differential band characteristic of the QAQB state. The negative and quite broad band observed in the first scan at 1738 cm1 is probably given by such an effect (the shift compared to the 1735 cm1 band could possibly arise from a broadening effect due to the rapidscan technique combined with the overlapping rising 1728 cm1 positive band, although some spectral shift induced by the presence of Cd2+ cannot be excluded). Interestingly, there is no evidence for the 1707 cm1 band in any of the spectra.5 This is quite surprising, because we could expect that if Cd2+ binding slows down the electron transfer reaction, it should also slow down the proton transfer reaction, making the observation of the 1707 cm1 intermediate more easy (as in the case of the low-temperature experiments shown before). The absence of the 1707 cm1 band could be explained by the fact that Cd2+ binding affects the proton pathway, hampering the protonation of the Asp or Glu side chain responsible for the 1707 cm1. The crystal structure of the RC shows that Asp-M17 and Asp-L210 are extremely close to the binding site of Cd2+ [39], suggesting that the presence of the metal could modify their pKa and therefore their availability to take part in the proton transfer mechanism. Furthermore, recent calculations have shown that the effect of pKa modification induced by the presence of Cd2+ is strong and affects several other protonable amino acid side chains including Asp-L213, ˚ ) from the binding which is located relatively far away (12 A 2+ site of Cd [40]. These pKa shifts are probably responsible for a slow proton transfer to the QB site which in turn slows down also the electron transfer reaction [40]. It is important to mention that preliminary rapid-scan experiments on Ni2+-treated RCs (not shown) show a similar effect (i.e. absence of the 1707 cm1 band in all the spectra). Ni2+ binds to the RC at the His-H126 and AspM17 residues, very close to the Cd2+ binding site [40]. It is difficult to identify which is the amino acid whose side chain is responsible for the 1707 cm1 band, as all residues involved in the proton transfer pathway are possibly interconnected so that the presence of a metal or the mutation of an amino acid can in principle affect other residues. In the previous paragraph we have suggested that the 1707 cm1 band could arise from the Asp-M17 side chain. Nevertheless, more detailed investigations on mutant RCs will be necessary in order to unambiguously identify the residue responsible of the transient 1707 cm1 band.
5
2+
Fig. 7. Rapid-scan FTIR spectra recorded after the 1st flash in Cd -treated wild-type RCs. T = 281 K.
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It should however be noted that, given the relatively high noise level, the presence of very small contributions of the 1707 cm1 band in the first spectrum cannot be totally excluded.
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4. Conclusions In this paper, we have shown that rapid-scan FTIR difference spectroscopy can be successfully used to investigate lightinduced reactions in bacterial RCs. From a methodological point of view, we have shown that quite simple strategies and/or experiment design can considerably increase the amount of information that can be obtained from spectral data. In particular, combination of experiments at different temperatures, in different media (H2O and D2O), on mutant RCs, or in presence of proton uptake inhibitors has allowed to extract from rapid-scan data new and interesting pieces of information concerning the proton and electron transfer reactions and the coupling between these two physicochemical processes. More precisely, we have shown that (1) the 1707 cm1 transient band observed after the 1st and the 2nd flashes is sensitive to temperature and to H/D exchange. This strongly suggests its identification with a Glu or Asp residue side chain transient protonation; (2) the QAQB ! QAQB electron transfer reaction in the AspM17 ! Asn mutant is slowed down by H/D exchange. This effect shows that in this mutant electron transfer is ratelimited by a process (involving proton movements in the protein) which is strongly influenced by H/D exchange; (3) consistent with previous UV–vis measurements, the QAQB ! QAQB electron transfer reaction is slowed down in presence of a proton uptake inhibitor (Cd2+); (4) in Cd2+-treated RCs the transient 1707 cm1 band is not observed, possibly because of a modification of the whole proton transfer pathway upon Cd2+ binding. Therefore this suggests its identification with a residue located on the proton transfer pathway. Acknowledgments The authors thank Prof. M.Y. Okamura and Prof. M.L. Paddock for the gift of the Asp-M17 ! Asn mutant RCs. A.M. thanks CEA/Saclay, the ‘‘Angelo Della Riccia’’ and ‘‘Guido Donegani’’ Foundations for partial financial support. References [1] P. Mitchell, J. Moyle, The role of ubiquinone and plastoquinone in chemiosmotic coupling between electron transfer and proton translocation, in: G. Lenaz (Ed.), Coenzyme Q, J Wiley & Sons, Chichester, 1985, p. 145. [2] J.P. Allen, G. Feher, T.O. Yeates, H. Komyia, D.C. Rees, Proc. Natl. Acad. Sci. U.S.A. 85 (1988) 8487. [3] U. Ermler, G. Fritzsch, S. Buchanan, H. Michel, Structure 2 (1994) 925. [4] G. Fritzsch, J. Koepke, R. Diem, A. Kuglstatter, L. Baciou, Acta Crystallogr. D: Biol. Crystallogr. 58 (2002) 1660. [5] M.Y. Okamura, M.L. Paddock, M.S. Graige, G. Feher, Biochim. Biophys. Acta: Bioenergetics 1458 (2000) 148. [6] M.L. Paddock, G. Feher, M.Y. Okamura, FEBS Lett. 555 (2003) 45. [7] M.S. Graige, G. Feher, M.Y. Okamura, Proc. Natl. Acad. Sci. U.S.A. 95 (1998) 11679. [8] M.H.B. Stowell, T.M. McPhillips, D.C. Rees, S.M. Soltis, E. Abresch, G. Feher, Science 276 (1997) 812.
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