Histidine is involved in coupling proton uptake to electron transfer in photosynthetic proteins

European Journal of Cell Biology 89 (2010) 983–989

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European Journal of Cell Biology journal homepage: www.elsevier.de/ejcb

Histidine is involved in coupling proton uptake to electron transfer in photosynthetic proteins Delphine Onidas a,b,1 , Joanna M. Stachnik c,1 , Sven Brucker c , Steffen Krätzig c , Klaus Gerwert c,∗ a b c

UFR Biomédicale, Université Paris Descartes, 45 rue des Saints-Pères, F-75006 Paris, France Université Paris-Sud/CNRS, Laboratoire de Chimie-Physique, UMR 8000, Orsay F-91405, France Lehrstuhl für Biophysik, ND 04/596, Ruhr-Universität Bochum, D-44780 Bochum, Germany

a r t i c l e

i n f o

Keywords: Photosynthesis Reaction center Electron transfer L210DN Histidine Fourier transform infrared Isotopic labeling Band assignment Protonated water

a b s t r a c t In photosynthesis, the central step in transforming light energy into chemical energy is the coupling of light-induced electron transfer to proton uptake and release. Despite intense investigations of different photosynthetic protein complexes, including the photosystem II (PS II) in plants and the reaction center (RC) in bacteria, the molecular details of this fundamental process remain incompletely understood. In the RC of Rhodobacter (Rb.) sphaeroides, fast formation of the charge separated state, P+ QA − , is followed by a slower electron transfer from the primary acceptor, QA , to the secondary acceptor, QB , and the uptake of a proton from the cytoplasm. The proton transfer to QB takes place via a protonated water chain. Mutation of the amino acid AspL210 to Asn (L210DN mutant) near the entry of the proton pathway can disturb this water chain and consequently slow down proton uptake. Time-resolved step-scan Fourier transform infrared (FTIR) measurements revealed an intermediate X in the QA − QB to QA QB − transition. The nature of this transition remains a matter of debate. In this study, we investigated the role of the iron–histidine complex located between QA and QB . We used time-resolved fast-scan FTIR spectroscopy to characterize the Rb. sphaeroides L210DN RC mutant marked with isotopically labeled histidine. FTIR marker bands of the intermediate X between 1120 cm−1 and 1050 cm−1 are assigned to histidine vibrations and indicate the protonation of a histidine, most likely HisL190, during the disappearance of the intermediate. Based on these results we propose a novel mechanism of the coupling of electron and proton transfer in photosynthesis. © 2010 Elsevier GmbH. All rights reserved.

Introduction In photosynthesis and respiration, a proton gradient is created across a membrane to establish the driving force for ATPases to synthesize ATP, the fuel of life. In photosynthesis, the proton gradient is formed by coupling light-induced electron transfer to proton uptake and release. Here, we investigated how this coupling is achieved at the atomic level. The coupling process takes place within a membrane protein complex, either in the reaction center (RC) in bacterial photosynthesis, or in photosystem II (PS II) in plant photosynthesis. The photosynthetic RC from the purple nonsulfur bacterium Rhodobacter (Rb.) sphaeroides was used in this study. This pigment-containing transmembrane protein complex of ∼100 kDa is structurally (Ermler et al., 1994) and mechanistically (Wraight, 2004) well characterized, and very similar, in both aspects, to the photosystem II in higher plants (Zouni et al., 2001).

∗ Corresponding author. Tel.: +49 234 3224461; fax: +49 234 3214238. E-mail address: [email protected] (K. Gerwert). 1 These authors contributed equally. 0171-9335/$ – see front matter © 2010 Elsevier GmbH. All rights reserved. doi:10.1016/j.ejcb.2010.08.007

Light induces electron transfer from the primary electron donor P, a dimer of bacteriochlorophyll a molecules, via the intermediate acceptors, bacteriochlorophyll a (BA ) and bacteriopheophytin a (HA ) to the primary electron acceptor, quinone QA , and subsequently, to the secondary electron acceptor, quinone QB (Fig. 1). The P+ QA − charge separation occurs within 200 ps (Holzapfel et al., 1990), whereas the electron transition from QA − to QB is slow (200 ␮s) (Okamura et al., 2000; Tiede et al., 1996; Graige et al., 1998; Li et al., 1998, 2000). The QA − to QB electron transition is coupled to proton uptake from the cytoplasm (Okamura et al., 2000; Paddock et al., 2003), and leads to the formation of quinol (QB H2 ), which is released from the RC. After the fast formation of the P+ QA − charge separated state, AspL210, an amino acid close to the entry of the proton uptake pathway, becomes protonated within 12 ␮s (Remy and Gerwert, 2003). Further proton transfer from AspL210 to QB takes place in a Grotthus-like proton transfer mechanism via a protonated water chain. This water chain is disturbed by mutation of AspL210 to Asn (L210DN mutant), as shown in the X-ray structural model of the L210DN mutant RC (Hermes et al., 2006b). The disturbed water chain explains the slower proton uptake in the L210DN

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Materials and methods Chemicals Histidine hydrochloride monohydrate and histidine were provided by Sigma and SERVA Electrophoresis GmbH, respectively. 13 C ,15 N 6 3 labeled histidines were purchased from Eurisotop (France). Both 13 C and 15 N were 98% enrichments. Growth and purification

Fig. 1. Structure of the Rhodobacter sphaeroides reaction center. Overall organization of the reaction center cofactors, represented with sticks. The cofactors include the special pair (P: green), accessory bacteriochlorophyll a (BA/B : green), bacteriopheophytin a (HA/B : cyan), and quinones (QA/B : blue). The carotenoid (sphaeroidenone) is not shown. The iron molecule is shown (orange sphere). The main chains L, M, and H are illustrated with gray cartoons. The cofactors are arranged around an axis of 2-fold pseudosymmetry, in two branches (indices A and B) that span the membrane. The route of electron transfer from P to QB along the A-branch is indicated with red arrows.

mutant compared to the wild type (WT) RC. The deprotonation of AspL210 is also rate-limiting for the QA − QB → QA QB − electron transition. A QB movement as proposed by Stowell et al. (1997), could be excluded as the rate-limiting step (Remy and Gerwert, 2003). Time-resolved step-scan Fourier transform infrared (FTIR) experiments revealed an intermediate X in the QA − QB → QA QB − transition (Remy and Gerwert, 2003). These results have been widely accepted. These data then led to the question: is an intermediary electron donor involved in the intermediate X? A potential donor is the nonheme iron located between QA and QB . However, experiments with Fe-specific time-resolved X-ray absorption spectroscopy (XAS) have shown no evidence for a full oxidation of the Fe upon electron transfer from QA − to QB (Hermes et al., 2006a). Therefore, the Fe2+ alone was excluded as an intermediary electron donor to QB . Alternatively, another candidate might be a larger complex that includes the iron coordinating histidines. In fact, the integrity of the RC is very sensitive to any mutation of these histidines (Chirino et al., 1994; Remy, 2002), which suggests that the histidines might be involved in the electron transfer from QA to QB . In this study, we probed the potential involvement of these histidines with time-resolved FTIR experiments on RCs marked noninvasively with isotopically labeled histidines. We characterized the L210DN RC mutant, because it is slower than the WT, and thus, it allows detection with the fast-scan FTIR technique. We assign the bands between 1120 cm−1 and 1050 cm−1 of intermediate X now to histidine vibrations. The bands indicate protonation of a histidine, most likely HisL190. Based on these results, we propose a novel mechanism for the coupling of electron and proton transfer in photosynthesis.

The Rb. sphaeroides L210DN RC mutant was constructed as described (Farchaus and Oesterhelt, 1989) and modified (Remy, 2002). Growth of the mutant cells was performed as described by Hermes et al. (2006b) and was adapted to the incorporation of labeled histidine, as follows. Mutant cells were grown in an optimized synthetic medium according to Raap et al. (1990) for 3 days in the dark under semi-anaerobic conditions. The optimized synthetic medium had the following composition: 1.4 mM nitrilotriacetic acid, 2.4 mM MgSO4 , 575 ␮M CaCl2 , 58 ␮M FeSO4 , 0.16 ␮M (NH4 )6 Mo7 O24 , 220 ␮M l-Asp, 132 ␮M l-Glu, 370 ␮M lIle, 440 ␮M l-Phe, 360 ␮M l-Pro, 320 ␮M l-Ser, 250 ␮M l-Gln, 300 ␮M l-Thr, 280 ␮M l-Trp, 700 ␮M l-Leu, 500 ␮M l-His, 190 ␮M l-Arg, 46 ␮M l-Cys, 400 ␮M l-Val, 630 ␮M l-Ala, 190 ␮M l-Met, 200 ␮M l-Tyr, 200 ␮M l-Asn, 160 ␮M l-Lys, 660 ␮M l-Gly, 13.6 mM K2 HPO4 , 9.6 mM KH2 PO4 , 10.4 ␮M nicotinic acid, 3 ␮M thiamine, 0.08 ␮M biotin, 13 ␮M NaEDTA, 29 ␮M ZnSO4 , 5 ␮M MnCl2 , 1.2 ␮M CuSO4 , 0.6 ␮M CoCl2 , 1.3 ␮M H3 BO3 , 20 mM dl-malic acid, adjusted to pH 6.9 with NH3 , 43 ␮M kanamycin, and 4.2 ␮M tetracycline. For metabolic labeling experiments, histidine was replaced by its 13 C ,15 N isotopically labeled variant. Further purification steps 6 3 were performed, according to the procedure described by Hermes et al. (2006b), but scaled down for a smaller quantity of protein. As a control, L210DN RC was expressed and purified under the same conditions, but with Rb. sphaeroides grown in medium that contained histidine in its natural form. Mass spectrometry To isolate RC proteins, we loaded 1.5 ␮g of labeled and unlabeled proteins onto denatured, discontinuous 15% SDS polyacrylamide gels (Laemmli, 1970) for electrophoresis. After electrophoresis, the proteins were visualized with Coomassie® brilliant blue G250 staining (Bradford, 1976) and the respective bands of the L, M, and H subunits were manually excised from the gel. The gel-embedded proteins were transferred into mini glass tubes. In-gel digestion, mass spectrometry (MS), and analysis of mass spectrometric data were performed as described by Warscheid et al. (2008). FTIR measurements Purified proteins were concentrated with Vivaspin 500 concentrators (MWCO 30 kDa) for a final concentration of 8 mg/mL. Samples for the FTIR measurements were prepared as described previously (Hermes et al., 2006b). A protein solution that contained approximately 150 ␮g of RC (∼30 ␮L) was pipetted onto a CaF2 window and concentrated under a gentle nitrogen stream to 1 ␮L. The QB -activity was checked by recording P+ -kinetics at 960 nm, and the result showed ∼60% activity. Fast-scan FTIR measurements were carried out on an IFS 88 or IFS 66v (Bruker) at 278 K as described previously (Hermes et al., 2006b). The reaction was started either by an excimer-pumped dye laser (532 nm, 30 ns, ∼5 mJ) or by a flash lamp-pumped Nd–YAG laser (Continuum, Minilite II, 532 nm, 3–5 ns, ∼25 mJ). Data from three samples were averaged for each measurement. P+ QA − /PQA and P+ QB − /PQB steady-state spectra were respectively recorded at

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223 K and 278 K on an IFS 88 (Bruker) with a Fiberoptic-Heim LQ 2600 lamp for illumination (Hermes et al., 2006b). Histidine IR spectra at different pH strengths were recorded at room temperature on the ␮-ATR (attenuated total reflection) setup (equipped with a diamond crystal) of an IFS 88 spectrometer. The spectrum of solvent (15 ␮L of H2 O; Tris buffer; or NaOH solution) was measured (256 scans) and subtracted from the sample (15 ␮L of His 100 mM) spectrum (256 scans) to eliminate the high background of H2 O absorption. Fitting procedure All data were analyzed with the Global fit method, including all wave numbers from 1900 cm−1 to 950 cm−1 (Hessling et al., 1993). Results and discussion Isotopic labeling of RC Isotopically labeled histidines were biosynthetically incorporated into the RC. The incorporation was determined by mass spectrometry. Fig. 2A shows the MS spectrum of the doubly charged tryptic peptide AAAGFHVSAGK, isolated from unlabeled RC L210DN. The monoisotopic peak was detected at m/z 508.31 and the natural isotopic peaks of the peptide were observed at higher m/z values; each showed a mass shift of m/z = 0.5. Isotopically labeled RC showed the expected 4.5 Da mass shift at m/z 512.81 (Fig. 2B). The difference between the monoisotopic peak intensities of the unlabeled and labeled AAAGFHVSAGK peptides indicated that isotopically labeled histidine was incorporated at an efficiency of 75%. The presence of unlabeled L210DN RC in isotopically labeled culture was due to the use of an inoculation volume that contained unlabeled histidine in PY medium. Furthermore, histidine is produced endogenously by the natural metabolism of Rb. sphaeroides. IR spectra of isotopically labeled histidines at different pHs We recorded the IR spectra of two unlabeled histidine model compounds (4-methylimidazole and imidazole) and of 13 C6 ,15 N3 labeled histidine at different pH strengths. The IR spectra of histidine and histidine model compounds have been previously published at different pHs (Hasegawa et al., 2000; Iwaki et al., 2005; Wolpert and Hellwig, 2006), but not with isotopic labeling. The imidazole group of histidine has two nitrogen atoms that can be protonated or deprotonated. At pH 4, the fully protonated imidazolium cation is formed (Fig. 3A). At pH 8 the neutral imidazole group adopts two tautomeric forms in which one of the two nitrogen atoms is protonated (Fig. 3B). The protonation state of the imidazole group can be identified with several marker bands in the IR spectrum. A useful marker band is the CN-stretching vibration band near 1100 cm−1 (Hasegawa et al., 2000). The fully protonated form of histidine appears as one significant band at (+) 1094 cm−1 (Fig. 3A, green line); in contrast, the singly protonated histidine appears as two characteristic bands at (+) 1106 cm−1 and (+) 1089 cm−1 (Fig. 3B, green line), which correspond to the two tautomeric forms. Those results agree closely with the histidine band assignments of Barth (2000) and Hasegawa et al. (2000). The 13 C ,15 N labeled histidine shows corresponding shifts at pH 8 and 6 3 pH 4, as shown in Fig. 3A and B. In the fully protonated histidine, the characteristic band at (+) 1094 cm−1 shifts by 21 cm−1 . The two intense bands of singly protonated histidine at (+) 1106 cm−1 and (+) 1089 cm−1 shift by 26 cm−1 and 18 cm−1 , respectively. In Fig. 3C, the IR difference spectrum of pH 8 minus pH 4 is shown. It reveals the marker bands for a protonation change of histidine. Both the unlabeled histidine and the 13 C6 ,15 N3 labeled histidine difference spectra are shown. The difference spectrum

Fig. 2. Mass spectra of the AAAGFHVSAGK peptide. (A, B) Mass spectra of the doubly charged tryptic peptide, AAAGFHVSAGK ([M+ 2H]2+ = 508.31 Da). Peptides were isolated from (A) unlabeled RC L210DN and (B) 13 C6 ,15 N3 histidine labeled RC L210DN. The observed mass shift of 4.5 Da between the unlabeled and labeled tryptic peptides shows the incorporation of 13 C6 ,15 N3 histidine into RC protein. On the basis of the peak intensities at m/z 508.31 and 512.81, the level of metabolic labeling of L210DN RC protein was determined to be 75%. (C) Structure of the 13 C6 ,15 N3 labeled histidine. Red stars correspond to 13 C and 15 N atoms.

of the native histidine shows three significant marker bands at (+) 1107 cm−1 , (−) 1097 cm−1 , and (+) 1088 cm−1 . Due to labeling, these bands shift by 25 cm−1 , 21 cm−1 , and 19 cm−1 , respectively. Assignment of histidine bands in the RC The P+ QA − X+ /PQA X and P+ QA − /PQA amplitude spectra (Hermes et al., 2006b) of the L210DN RC mutant in the 1120–1050 cm−1 spectral region (Fig. 4A) were examined to identify bands that correspond to the X+ /X transition i.e., bands that are present in the P+ QA − X+ /PQA X spectrum, but missing in the P+ QA − /PQA spectrum. This comparison reveals three bands attributed to X (blue in Fig. 4A) at (+) 1110 cm−1 , (−) 1100 cm−1 and (+) 1068 cm−1 . In addition, a fraction of the band at (+) 1094 cm−1 could be considered an X band, because this band was also present in the QA − X+ /QA X amplitude spectrum (i.e., without the P+ /P contribution; Hermes et al., 2006b), and might be partially masked by P+ /P contributions.

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arising from histidine should be shifted in amplitude spectra of labeled protein. Fig. 4B shows the P+ QA − X+ /PQA X amplitude spectra of both the unlabeled L210DN mutant and the 13 C6 ,15 N3 labeled histidine L210DN RC protein in the 1120–1050 cm−1 region. The bands at (+) 1110 cm−1 , (−) 1100 cm−1 , and (+) 1094 cm−1 that were assigned to X (blue; compare with Fig. 4A) had shifted by approximately 8–13 cm−1 with the isotopic labeling of histidine. Therefore, for the first time, these could be clearly assigned to histidine vibrations. However, we assume that isolated amino acids in solution must exhibit different spectroscopic behavior than those within the interior of a hydrophobic protein. IR vibrations are very sensitive to the microenvironment. Therefore, we would expect slightly different band positions and frequency shifts for the histidine bands within the RC (Fig. 4B) compared to the difference spectra of isolated histidine (Fig. 3C). Additionally, the three spectral features observed in Fig. 4B are consistent with two other reports of similar features in the same spectral region; one studied the RC with 13 C histidine labeling (Breton et al., 2001) and the other studied PS II with 15 N histidine labeling (Hienerwadel and Berthomieu, 1995); this agreement further supports our assignment. Our control experiments with divalent metal ions show no deviations on the histidine vibrations between 1120 cm−1 and 1050 cm−1 . The histidines HisH126 and HisH128 at the proton entrance can be excluded to cause the absorbance changes in this region. Protonation state change of histidine In Fig. 4C, the P+ QA − X+ /PQA X amplitude spectrum of L210DN RC mutant is compared to the IR marker bands of protonation change in histidine (Fig. 3C) to determine whether a change in the protonation state of histidine had occurred during the QA − X+ to QA X transition. These spectra reveal the same marker bands in isolated histidine (green) and in the RC (blue), but slightly shifted in the RC protein interior, as expected. In particular, the two features (+) 1110 cm−1 /(−) 1100 cm−1 and (+) 1107 cm−1 /(−) 1097 cm−1 are nearly an exact match in frequency and band intensity. From this comparison, we propose that, in the P+ QA − X+ /PQA X amplitude spectrum, the positive bands at 1110 cm−1 and 1094 cm−1 correspond to a singly protonated histidine, and the negative band at 1100 cm−1 is assigned to a fully protonated histidine. An amplitude spectrum describes the absorbance changes involved in the respective transition: negative bands correspond to new species that appear during the transition, and positive bands represent the species that disappear. Therefore, the result depicted in Fig. 4C indicates a protonation of histidine during the QA − X+ to QA X transition. Fig. 3. Histidine bands (1120–1050 cm−1 ). Steady-state ␮-ATR (attenuated total reflection) IR spectra of histidine. Unlabeled histidine is shown with a green line; 13 C6 ,15 N3 labeled histidine is shown with a black line. Bands are labeled with their respective wave numbers. Bands of interest are highlighted in the respective color. Band shifts are indicated with red arrows and labeled with the respective difference in wave number (cm−1 ). (A) Histidine at pH 4 (fully protonated). (B) Histidine at pH 8 (singly protonated). (C) Difference spectrum of histidine (pH 8 minus pH 4).

In this study, we measured the FTIR amplitude spectra of the L210DN RC mutant labeled with 13 C6 ,15 N3 histidine, in order to clearly assign the appropriate bands of intermediate X to histidine vibrations. As a control, amplitude spectra were also recorded of unlabeled L210DN RC mutant protein, expressed and purified following exactly the same protocol as that applied to the labeled protein. The amplitude spectra of the unlabeled L210DN mutant (not shown) are similar to the amplitude spectra of the unlabeled L210DN mutant published previously (Hermes et al., 2006b). Compared to amplitude spectra of unlabeled protein only the bands

Novel mechanism for the coupling of electron transfer to proton uptake Based on these results, we propose a modification to the mechanism for the coupling of light-induced electron transfer to proton uptake, as described by Remy and Gerwert (2003). The original mechanism and the modification are illustrated in Fig. 5. In both mechanisms, the absorption of one photon induces a very fast electron transfer (200 ps) from P to QA (step 1, white arrow). A hydrogen-bonded network exists between QA and the cytoplasm via the Fe–histidine complex and the water chain from QB to AspL210 (gray dashed lines). Also, GluL212, GluH173, and GluH122 might be involved. Due to the long-range electrostatic effects via this hydrogen-bonded network, the reduction of QA appears to induce the uptake of a proton from the cytoplasm. The proton compensates the negative charge within the hydrophobic protein interior and facilitates the electron transfer from QA to QB . This

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Fig. 4. Assignment of histidine bands in RC L210DN (1120–1050 cm−1 ). Bands are labeled with their respective wave numbers. Bands of interest are highlighted in the respective color. (A) Comparison of the P+ QA − X+ /PQA X (blue line) and P+ QA − /PQA (black line) amplitude spectra of the L210DN mutant. (B) Comparison of the P+ QA − X+ /PQA X amplitude spectra of unlabeled (blue line) and isotopically labeled (red line) L210DN mutant. (C) Comparison of the P+ QA − X+ /PQA X amplitude spectrum of unlabeled L210DN mutant (blue line) with the difference spectrum of histidine (green line, pH 8 minus pH 4).

explains why the last electron transfer step from QA to QB is relatively slow, compared to the P to QA transition. This proton is taken up from the cytoplasm by AspL210 in 12 ␮s (step 2; yellow arrows), as shown by Remy and Gerwert (2003). Based on the present study, we propose a deprotonation of a histidine in 150 ␮s, most likely HisL190 in the P+ QA − X/P+ QA − X+ transition (step 3). Also, a carbonyl band of QB appears at 1479 cm−1 in step 3 (Remy and Gerwert, 2003). The appearance of this band previously led to the conclusion that QB was reduced in 150 ␮s, because this band was commonly taken as a QB − marker band (Breton et al., 1995; Brudler et al., 1995; Nabedryk et al., 1995). Based on that interpre-

tation an additional intermediary electron donor X between QA and QB was proposed, which first reduced QB and then became reoxidized by QA . This proposal is currently controversial, because both quinones would have to carry a negative charge at the same time, and the Fe has been excluded as an intermediary electron donor. However, the results of the present study offer an alternative explanation for the downshift of the carbonyl vibration from 1660 cm−1 to 1479 cm−1 . The downshift indicates a decrease of the C O bond order. We suggest that this might not be due to the formation of a C–O− bond of the QB semiquinone. Instead, a partial protonation of the 4C O carbonyl group of QB may also result in a decrease of the

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Fig. 5. Proposed mechanism for the electron and proton transfers in bacterial RC. Structural details of the wild type RC (2J8C; Koepke et al., 2007). (A) Step 1, white arrow: reduction of QA in 200 ps; step 2, yellow arrows: protonation of AspL210 in 12 ␮s; step 3, orange arrow: deprotonation of HisL190, protonation of putative flexible water molecule (black sphere), and decrease of the C O bond order of QB in 150 ␮s; step 4, red arrows: coupled GluL212 protonation, QB reduction, and QA oxidation in 1.1 ms. (B) Detailed structure of the shared proton between the QB C O bond and histidine.

C O bond order and that might shift the band from 1660 cm−1 to 1479 cm−1 . The deprotonation of HisL190 might provide a proton to the water molecule (black sphere) between QB and HisL190, as illustrated in Fig. 5, step 3 (orange arrow). This water molecule was not resolved in the recently published structure, 2J8C (Koepke et al., 2007). However, those authors suggested that there was enough space for a flexible water molecule close to QB . This additional mobile water molecule placed between the Fe–histidine complex and QB at hydrogen bond distances from amino acids and cofactors might supply the missing link in the hydrogen-bonded network and complete the connections between the surface, GluM234, and the carbonyl oxygen of QB . Thus, we can summarize step 3 occurring in 150 ␮s, as follows: a histidine protonates the flexible water (black sphere) located between HisL190 and QB . This protonated water interacts with the C O group of QB , leading to a bond order decrease of the C O bond. Thus, the QB site is partially solvated for the electron transfer from QA , and the protonated H-bonded network, which ranges from QB to AspL210, is prepared for proton transfer. In step 4, which takes place in 1.1 ms, the electron is transferred from QA − to QB , and the histidine is reprotonated. This event is coupled to the proton transfer from AspL210 to GluL212, which occurs via a chain of protonated water molecules in a Grotthuslike mechanism (Hermes et al., 2006b). Step 4 is consistent with results of Hienerwadel et al. (1995) who proposed that protonation of GluL212 along with the protonation or deprotonation of other ionizable residues provides charge compensation for QB − . The protonation of AspL210 might trigger the coupled electron–proton transfer. Indeed, when AspL210 is protonated, proton and electron transfers can occur; conversely, when AspL210 is deprotonated (i.e., L210DN mutant), the gate is closed and no electron–proton event can occur in 1.1 ms. Due to the protonation of GluL212 in step 4, QB is solvated by the positive charges of the protonated GluL212, which facilitates the transfer of the electron from QA to QB . Further studies with QM/MM simulations should consider these results and endeavor to provide a more quantitative description of these

events. In particular, it would be experimentally challenging to calculate the 4C O vibration of QB in this scenario and to calculate and measure the continuum bands, which would indicate the presence of a protonated water chain. Conclusion In this study, we have shown that a histidine, most likely HisL190, is successively deprotonated and reprotonated in the intermediate X during the QA − QB /QA QB − transition. The proton appears to be shared between HisL190, a water molecule, and QB . This induces protonation of GluL212 via a protonated water chain and creates the electrostatic environment around the QB site necessary for the final electron transfer step from QA to QB . Acknowledgments This work was supported by the SFB480-C3 grant from the Deutsche Forschungsgemeinschaft. D.O. gratefully acknowledges a fellowship from the Alexander von Humboldt Foundation. J.M.S. gratefully acknowledges a fellowship from the Ruhr University Research School and the Günther and Wilhelm Esser Foundation. The authors wish to thank Yvonne Fretter for technical assistance in protein production and purification. Thanks also to Bettina Warscheid and Heike Piechura for performing the mass spectrometry. References Barth, A., 2000. The infrared absorption of amino acid side chains. Prog. Biophys. Mol. Biol. 74, 141–173. Bradford, M.M., 1976. A Rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein–dye binding. Anal. Biochem. 72, 248–254. Breton, J., Boullais, C., Berger, G., Mioskowski, C., Nabedryk, E., 1995. Binding sites of quinones in photosynthetic bacterial reaction centers investigated by

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