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

Contents lists available at ScienceDirect

European Journal of Cell Biology

journa l homepage: www.e lsev ier .de /e jcb

istidine is involved in coupling proton uptake to electron transfer inhotosynthetic proteins

elphine Onidasa,b,1, Joanna M. Stachnikc,1, Sven Bruckerc, Steffen Krätzigc, Klaus Gerwertc,∗

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

r t i c l e i n f o

eywords:hotosynthesiseaction centerlectron transfer210DNistidineourier transform infraredsotopic labelingand assignmentrotonated water

a b s t r a c t

In photosynthesis, the central step in transforming light energy into chemical energy is the coupling oflight-induced electron transfer to proton uptake and release. Despite intense investigations of differentphotosynthetic 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. Inthe RC of Rhodobacter (Rb.) sphaeroides, fast formation of the charge separated state, P+QA

−, is followed bya slower electron transfer from the primary acceptor, QA, to the secondary acceptor, QB, and the uptake ofa proton from the cytoplasm. The proton transfer to QB takes place via a protonated water chain. Mutationof the amino acid AspL210 to Asn (L210DN mutant) near the entry of the proton pathway can disturbthis water chain and consequently slow down proton uptake. Time-resolved step-scan Fourier transforminfrared (FTIR) measurements revealed an intermediate X in the QA

−QB to QAQB− transition. The nature

of this transition remains a matter of debate. In this study, we investigated the role of the iron–histidinecomplex located between QA and QB. We used time-resolved fast-scan FTIR spectroscopy to characterizethe Rb. sphaeroides L210DN RC mutant marked with isotopically labeled histidine. FTIR marker bandsof the intermediate X between 1120 cm−1 and 1050 cm−1 are assigned to histidine vibrations and indi-cate 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.

ntroduction

In photosynthesis and respiration, a proton gradient is cre-ted across a membrane to establish the driving force for ATPaseso synthesize ATP, the fuel of life. In photosynthesis, the protonradient is formed by coupling light-induced electron transfer toroton uptake and release. Here, we investigated how this cou-ling is achieved at the atomic level. The coupling process takeslace within a membrane protein complex, either in the reactionenter (RC) in bacterial photosynthesis, or in photosystem II (PSI) in plant photosynthesis. The photosynthetic RC from the purpleonsulfur bacterium Rhodobacter (Rb.) sphaeroides was used in this

tudy. This pigment-containing transmembrane protein complexf ∼100 kDa is structurally (Ermler et al., 1994) and mechanisti-ally (Wraight, 2004) well characterized, and very similar, in bothspects, to the photosystem II in higher plants (Zouni et al., 2001).

∗ Corresponding author. Tel.: +49 234 3224461; fax: +49 234 3214238.E-mail address: gerwert@bph.rub.de (K. Gerwert).

1 These authors contributed equally.

171-9335/$ – see front matter © 2010 Elsevier GmbH. All rights reserved.oi:10.1016/j.ejcb.2010.08.007

© 2010 Elsevier GmbH. All rights reserved.

Light induces electron transfer from the primary electron donorP, a dimer of bacteriochlorophyll a molecules, via the intermedi-ate acceptors, bacteriochlorophyll a (BA) and bacteriopheophytina (HA) to the primary electron acceptor, quinone QA, and subse-quently, to the secondary electron acceptor, quinone QB (Fig. 1).The P+QA

− charge separation occurs within 200 ps (Holzapfel etal., 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 toproton uptake from the cytoplasm (Okamura et al., 2000; Paddocket al., 2003), and leads to the formation of quinol (QBH2), which isreleased 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 uptakepathway, becomes protonated within 12 �s (Remy and Gerwert,2003). Further proton transfer from AspL210 to QB takes place in aGrotthus-like proton transfer mechanism via a protonated water

chain. This water chain is disturbed by mutation of AspL210 toAsn (L210DN mutant), as shown in the X-ray structural modelof the L210DN mutant RC (Hermes et al., 2006b). The disturbedwater chain explains the slower proton uptake in the L210DN

984 D. Onidas et al. / European Journal of C

Fig. 1. Structure of the Rhodobacter sphaeroides reaction center. Overall organizationof the reaction center cofactors, represented with sticks. The cofactors include thespecial pair (P: green), accessory bacteriochlorophyll a (BA/B: green), bacteriopheo-phytin a (HA/B: cyan), and quinones (QA/B: blue). The carotenoid (sphaeroidenone) isnot shown. The iron molecule is shown (orange sphere). The main chains L, M, andH are illustrated with gray cartoons. The cofactors are arranged around an axis of2-fold pseudosymmetry, in two branches (indices A and B) that span the membrane.Ta

motc2etw

dhFhtttto(ttwwRdbtmmt

2006b). The reaction was started either by an excimer-pumped dyelaser (532 nm, 30 ns, ∼5 mJ) or by a flash lamp-pumped Nd–YAG

he route of electron transfer from P to QB along the A-branch is indicated with redrrows.

utant compared to the wild type (WT) RC. The deprotonationf AspL210 is also rate-limiting for the QA

−QB → QAQB− electron

ransition. A QB movement as proposed by Stowell et al. (1997),ould be excluded as the rate-limiting step (Remy and Gerwert,003). Time-resolved step-scan Fourier transform infrared (FTIR)xperiments revealed an intermediate X in the QA

−QB → QAQB−

ransition (Remy and Gerwert, 2003). These results have beenidely accepted.

These data then led to the question: is an intermediary electrononor involved in the intermediate X? A potential donor is the non-eme iron located between QA and QB. However, experiments withe-specific time-resolved X-ray absorption spectroscopy (XAS)ave shown no evidence for a full oxidation of the Fe upon elec-ron transfer from QA

− to QB (Hermes et al., 2006a). Therefore,he Fe2+ alone was excluded as an intermediary electron donoro QB. Alternatively, another candidate might be a larger complexhat includes the iron coordinating histidines. In fact, the integrityf the RC is very sensitive to any mutation of these histidinesChirino et al., 1994; Remy, 2002), which suggests that the his-idines might be involved in the electron transfer from QA to QB. Inhis study, we probed the potential involvement of these histidinesith time-resolved FTIR experiments on RCs marked noninvasivelyith isotopically labeled histidines. We characterized the L210DNC mutant, because it is slower than the WT, and thus, it allowsetection with the fast-scan FTIR technique. We assign the bandsetween 1120 cm−1 and 1050 cm−1 of intermediate X now to his-idine vibrations. The bands indicate protonation of a histidine,

ost likely HisL190. Based on these results, we propose a novelechanism for the coupling of electron and proton transfer in pho-

osynthesis.

ell Biology 89 (2010) 983–989

Materials and methods

Chemicals

Histidine hydrochloride monohydrate and histidine were pro-vided by Sigma and SERVA Electrophoresis GmbH, respectively.13C6,15N3 labeled histidines were purchased from Eurisotop(France). Both 13C and 15N were 98% enrichments.

Growth and purification

The Rb. sphaeroides L210DN RC mutant was constructed asdescribed (Farchaus and Oesterhelt, 1989) and modified (Remy,2002). Growth of the mutant cells was performed as describedby Hermes et al. (2006b) and was adapted to the incorporationof labeled histidine, as follows. Mutant cells were grown in anoptimized synthetic medium according to Raap et al. (1990) for3 days in the dark under semi-anaerobic conditions. The opti-mized synthetic medium had the following composition: 1.4 mMnitrilotriacetic acid, 2.4 mM MgSO4, 575 �M CaCl2, 58 �M FeSO4,0.16 �M (NH4)6Mo7O24, 220 �M l-Asp, 132 �M l-Glu, 370 �M l-Ile, 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 �Ml-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 mMK2HPO4, 9.6 mM KH2PO4, 10.4 �M nicotinic acid, 3 �M thiamine,0.08 �M biotin, 13 �M NaEDTA, 29 �M ZnSO4, 5 �M MnCl2, 1.2 �MCuSO4, 0.6 �M CoCl2, 1.3 �M H3BO3, 20 mMdl-malic acid, adjustedto pH 6.9 with NH3, 43 �M kanamycin, and 4.2 �M tetracycline.For metabolic labeling experiments, histidine was replaced by its13C6,15N3 isotopically labeled variant. Further purification stepswere performed, according to the procedure described by Hermeset al. (2006b), but scaled down for a smaller quantity of protein.As a control, L210DN RC was expressed and purified under thesame conditions, but with Rb. sphaeroides grown in medium thatcontained histidine in its natural form.

Mass spectrometry

To isolate RC proteins, we loaded 1.5 �g of labeled and unlabeledproteins onto denatured, discontinuous 15% SDS polyacrylamidegels (Laemmli, 1970) for electrophoresis. After electrophoresis,the proteins were visualized with Coomassie® brilliant blue G250staining (Bradford, 1976) and the respective bands of the L, M, andH subunits were manually excised from the gel. The gel-embeddedproteins were transferred into mini glass tubes. In-gel digestion,mass spectrometry (MS), and analysis of mass spectrometric datawere performed as described by Warscheid et al. (2008).

FTIR measurements

Purified proteins were concentrated with Vivaspin 500 con-centrators (MWCO 30 kDa) for a final concentration of 8 mg/mL.Samples for the FTIR measurements were prepared as describedpreviously (Hermes et al., 2006b). A protein solution that containedapproximately 150 �g of RC (∼30 �L) was pipetted onto a CaF2 win-dow and concentrated under a gentle nitrogen stream to 1 �L. TheQB-activity was checked by recording P+-kinetics at 960 nm, andthe result showed ∼60% activity.

Fast-scan FTIR measurements were carried out on an IFS 88 orIFS 66v (Bruker) at 278 K as described previously (Hermes et al.,

laser (Continuum, Minilite II, 532 nm, 3–5 ns, ∼25 mJ). Data fromthree samples were averaged for each measurement. P+QA

−/PQAand P+QB

−/PQB steady-state spectra were respectively recorded at

al of Cell Biology 89 (2010) 983–989 985

22

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F

w

R

I

rscLnvRTutTwbe

I

clhp2icizntivp1h(tb1

pti(

shh

Fig. 2. Mass spectra of the AAAGFHVSAGK peptide. (A, B) Mass spectra of the doublycharged tryptic peptide, AAAGFHVSAGK ([M+2H]2+ = 508.31 Da). Peptides were iso-lated from (A) unlabeled RC L210DN and (B) 13C6,15N3 histidine labeled RC L210DN.The observed mass shift of 4.5 Da between the unlabeled and labeled tryptic pep-tides shows the incorporation of 13C ,15N histidine into RC protein. On the basis

D. Onidas et al. / European Journ

23 K and 278 K on an IFS 88 (Bruker) with a Fiberoptic-Heim LQ600 lamp for illumination (Hermes et al., 2006b).

Histidine IR spectra at different pH strengths were recorded atoom temperature on the �-ATR (attenuated total reflection) set-p (equipped with a diamond crystal) of an IFS 88 spectrometer.he spectrum of solvent (15 �L of H2O; Tris buffer; or NaOH solu-ion) was measured (256 scans) and subtracted from the sample15 �L of His 100 mM) spectrum (256 scans) to eliminate the highackground of H2O absorption.

itting procedure

All data were analyzed with the Global fit method, including allave numbers from 1900 cm−1 to 950 cm−1 (Hessling et al., 1993).

esults and discussion

sotopic labeling of RC

Isotopically labeled histidines were biosynthetically incorpo-ated into the RC. The incorporation was determined by masspectrometry. Fig. 2A shows the MS spectrum of the doublyharged tryptic peptide AAAGFHVSAGK, isolated from unlabeled RC210DN. The monoisotopic peak was detected at m/z 508.31 and theatural isotopic peaks of the peptide were observed at higher m/zalues; each showed a mass shift of m/z = 0.5. Isotopically labeledC showed the expected 4.5 Da mass shift at m/z 512.81 (Fig. 2B).he difference between the monoisotopic peak intensities of thenlabeled and labeled AAAGFHVSAGK peptides indicated that iso-opically labeled histidine was incorporated at an efficiency of 75%.he presence of unlabeled L210DN RC in isotopically labeled cultureas due to the use of an inoculation volume that contained unla-

eled histidine in PY medium. Furthermore, histidine is producedndogenously by the natural metabolism of Rb. sphaeroides.

R spectra of isotopically labeled histidines at different pHs

We recorded the IR spectra of two unlabeled histidine modelompounds (4-methylimidazole and imidazole) and of 13C6,15N3abeled histidine at different pH strengths. The IR spectra ofistidine and histidine model compounds have been previouslyublished at different pHs (Hasegawa et al., 2000; Iwaki et al.,005; Wolpert and Hellwig, 2006), but not with isotopic label-

ng. The imidazole group of histidine has two nitrogen atoms thatan be protonated or deprotonated. At pH 4, the fully protonatedmidazolium cation is formed (Fig. 3A). At pH 8 the neutral imida-ole group adopts two tautomeric forms in which one of the twoitrogen atoms is protonated (Fig. 3B). The protonation state ofhe imidazole group can be identified with several marker bandsn the IR spectrum. A useful marker band is the CN-stretchingibration band near 1100 cm−1 (Hasegawa et al., 2000). The fullyrotonated form of histidine appears as one significant band at (+)094 cm−1 (Fig. 3A, green line); in contrast, the singly protonatedistidine appears as two characteristic bands at (+) 1106 cm−1 and+) 1089 cm−1 (Fig. 3B, green line), which correspond to the twoautomeric forms. Those results agree closely with the histidineand assignments of Barth (2000) and Hasegawa et al. (2000). The3C6,15N3 labeled histidine shows corresponding shifts at pH 8 andH 4, as shown in Fig. 3A and B. In the fully protonated histidine,he characteristic band at (+) 1094 cm−1 shifts by 21 cm−1. The twontense 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 ishown. It reveals the marker bands for a protonation change ofistidine. Both the unlabeled histidine and the 13C6,15N3 labeledistidine difference spectra are shown. The difference spectrum

6 3

of the peak intensities at m/z 508.31 and 512.81, the level of metabolic labeling ofL210DN RC protein was determined to be 75%. (C) Structure of the 13C6,15N3 labeledhistidine. Red stars correspond to 13C and 15N 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+/PQAX and P+QA

−/PQA amplitude spectra (Hermeset al., 2006b) of the L210DN RC mutant in the 1120–1050 cm−1

spectral region (Fig. 4A) were examined to identify bands that cor-respond to the X+/X transition i.e., bands that are present in theP+QA

−X+/PQAX 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−1and (+) 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+/QAX amplitudespectrum (i.e., without the P+/P contribution; Hermes et al., 2006b),and might be partially masked by P+/P contributions.

986 D. Onidas et al. / European Journal of C

Fig. 3. Histidine bands (1120–1050 cm−1). Steady-state �-ATR (attenuated totalreflection) IR spectra of histidine. Unlabeled histidine is shown with a green line;13C6,15N3 labeled histidine is shown with a black line. Bands are labeled with theirrespective wave numbers. Bands of interest are highlighted in the respective color.Bi8

Lcvulp(Lp

might be involved. Due to the long-range electrostatic effects via

and shifts are indicated with red arrows and labeled with the respective differencen wave number (cm−1). (A) Histidine at pH 4 (fully protonated). (B) Histidine at pH(singly protonated). (C) Difference spectrum of histidine (pH 8 minus pH 4).

In this study, we measured the FTIR amplitude spectra of the210DN RC mutant labeled with 13C6,15N3 histidine, in order tolearly assign the appropriate bands of intermediate X to histidineibrations. As a control, amplitude spectra were also recorded ofnlabeled L210DN RC mutant protein, expressed and purified fol-

owing exactly the same protocol as that applied to the labeled

rotein. The amplitude spectra of the unlabeled L210DN mutantnot shown) are similar to the amplitude spectra of the unlabeled210DN mutant published previously (Hermes et al., 2006b). Com-ared to amplitude spectra of unlabeled protein only the bands

ell Biology 89 (2010) 983–989

arising from histidine should be shifted in amplitude spectra oflabeled protein.

Fig. 4B shows the P+QA−X+/PQAX amplitude spectra of both

the unlabeled L210DN mutant and the 13C6,15N3 labeled histidineL210DN RC protein in the 1120–1050 cm−1 region. The bands at (+)1110 cm−1, (−) 1100 cm−1, and (+) 1094 cm−1 that were assignedto X (blue; compare with Fig. 4A) had shifted by approximately8–13 cm−1 with the isotopic labeling of histidine. Therefore, for thefirst time, these could be clearly assigned to histidine vibrations.

However, we assume that isolated amino acids in solution mustexhibit different spectroscopic behavior than those within the inte-rior of a hydrophobic protein. IR vibrations are very sensitive to themicroenvironment. Therefore, we would expect slightly differentband positions and frequency shifts for the histidine bands withinthe RC (Fig. 4B) compared to the difference spectra of isolated histi-dine (Fig. 3C). Additionally, the three spectral features observed inFig. 4B are consistent with two other reports of similar featuresin the same spectral region; one studied the RC with 13C histi-dine labeling (Breton et al., 2001) and the other studied PS II with15N histidine labeling (Hienerwadel and Berthomieu, 1995); thisagreement further supports our assignment.

Our control experiments with divalent metal ions show nodeviations on the histidine vibrations between 1120 cm−1 and1050 cm−1. The histidines HisH126 and HisH128 at the protonentrance can be excluded to cause the absorbance changes in thisregion.

Protonation state change of histidine

In Fig. 4C, the P+QA−X+/PQAX amplitude spectrum of L210DN RC

mutant is compared to the IR marker bands of protonation changein histidine (Fig. 3C) to determine whether a change in the pro-tonation state of histidine had occurred during the QA

−X+ to QAXtransition. These spectra reveal the same marker bands in isolatedhistidine (green) and in the RC (blue), but slightly shifted in theRC 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. Fromthis comparison, we propose that, in the P+QA

−X+/PQAX amplitudespectrum, the positive bands at 1110 cm−1 and 1094 cm−1 corre-spond to a singly protonated histidine, and the negative band at1100 cm−1 is assigned to a fully protonated histidine.

An amplitude spectrum describes the absorbance changesinvolved in the respective transition: negative bands correspond tonew species that appear during the transition, and positive bandsrepresent the species that disappear. Therefore, the result depictedin Fig. 4C indicates a protonation of histidine during the QA

−X+ toQAX transition.

Novel mechanism for the coupling of electron transfer to protonuptake

Based on these results, we propose a modification to the mech-anism for the coupling of light-induced electron transfer to protonuptake, as described by Remy and Gerwert (2003). The originalmechanism and the modification are illustrated in Fig. 5. In bothmechanisms, the absorption of one photon induces a very fastelectron transfer (200 ps) from P to QA (step 1, white arrow). Ahydrogen-bonded network exists between QA and the cytoplasmvia the Fe–histidine complex and the water chain from QB toAspL210 (gray dashed lines). Also, GluL212, GluH173, and GluH122

this hydrogen-bonded network, the reduction of QA appears toinduce the uptake of a proton from the cytoplasm. The protoncompensates the negative charge within the hydrophobic proteininterior and facilitates the electron transfer from QA to QB. This

D. Onidas et al. / European Journal of Cell Biology 89 (2010) 983–989 987

F e laber lack lia mutam s pH 4

eatlpmAatwB

ig. 4. Assignment of histidine bands in RC L210DN (1120–1050 cm−1). Bands arespective color. (A) Comparison of the P+QA

−X+/PQAX (blue line) and P+QA−/PQA (b

mplitude spectra of unlabeled (blue line) and isotopically labeled (red line) L210DNutant (blue line) with the difference spectrum of histidine (green line, pH 8 minu

xplains why the last electron transfer step from QA to QB is rel-tively slow, compared to the P to QA transition. This proton isaken up from the cytoplasm by AspL210 in 12 �s (step 2; yel-ow arrows), as shown by Remy and Gerwert (2003). Based on theresent study, we propose a deprotonation of a histidine in 150 �s,ost likely HisL190 in the P+QA

−X/P+QA−X+ transition (step 3).

−1

lso, a carbonyl band of QB appears at 1479 cm in step 3 (Remynd Gerwert, 2003). The appearance of this band previously led tohe conclusion that QB was reduced in 150 �s, because this bandas commonly taken as a QB

− marker band (Breton et al., 1995;rudler et al., 1995; Nabedryk et al., 1995). Based on that interpre-

led with their respective wave numbers. Bands of interest are highlighted in thene) amplitude spectra of the L210DN mutant. (B) Comparison of the P+QA

−X+/PQAXnt. (C) Comparison of the P+QA

−X+/PQAX amplitude spectrum of unlabeled L210DN).

tation an additional intermediary electron donor X between QA andQB was proposed, which first reduced QB and then became reoxi-dized by QA. This proposal is currently controversial, because bothquinones 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 expla-

−1

nation for the downshift of the carbonyl vibration from 1660 cmto 1479 cm−1. The downshift indicates a decrease of the C O bondorder. We suggest that this might not be due to the formation of aC–O− bond of the QB semiquinone. Instead, a partial protonation ofthe 4C O carbonyl group of QB may also result in a decrease of the

988 D. Onidas et al. / European Journal of Cell Biology 89 (2010) 983–989

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:r stepm , red aD

C1tin2smamatr(wdfntfiolrtopttLcQwsa

eduction of QA in 200 ps; step 2, yellow arrows: protonation of AspL210 in 12 �s;olecule (black sphere), and decrease of the C O bond order of QB in 150 �s; step 4etailed structure of the shared proton between the QB C O bond and histidine.

O bond order and that might shift the band from 1660 cm−1 to479 cm−1. The deprotonation of HisL190 might provide a protono the water molecule (black sphere) between QB and HisL190, asllustrated in Fig. 5, step 3 (orange arrow). This water molecule wasot resolved in the recently published structure, 2J8C (Koepke et al.,007). However, those authors suggested that there was enoughpace for a flexible water molecule close to QB. This additionalobile water molecule placed between the Fe–histidine complex

nd QB at hydrogen bond distances from amino acids and cofactorsight supply the missing link in the hydrogen-bonded network

nd complete the connections between the surface, GluM234, andhe carbonyl oxygen of QB. Thus, we can summarize step 3 occur-ing in 150 �s, as follows: a histidine protonates the flexible waterblack sphere) located between HisL190 and QB. This protonatedater interacts with the C O group of QB, leading to a bond orderecrease of the C O bond. Thus, the QB site is partially solvatedor the electron transfer from QA, and the protonated H-bondedetwork, which ranges from QB to AspL210, is prepared for protonransfer. In step 4, which takes place in 1.1 ms, the electron is trans-erred from QA

− to QB, and the histidine is reprotonated. This events coupled to the proton transfer from AspL210 to GluL212, whichccurs via a chain of protonated water molecules in a Grotthus-ike mechanism (Hermes et al., 2006b). Step 4 is consistent withesults of Hienerwadel et al. (1995) who proposed that protona-ion of GluL212 along with the protonation or deprotonation ofther ionizable residues provides charge compensation for QB

−. Therotonation of AspL210 might trigger the coupled electron–protonransfer. Indeed, when AspL210 is protonated, proton and electronransfers can occur; conversely, when AspL210 is deprotonated (i.e.,210DN mutant), the gate is closed and no electron–proton event

an occur in 1.1 ms. Due to the protonation of GluL212 in step 4,B is solvated by the positive charges of the protonated GluL212,hich facilitates the transfer of the electron from QA to QB. Further

tudies with QM/MM simulations should consider these resultsnd endeavor to provide a more quantitative description of these

3, orange arrow: deprotonation of HisL190, protonation of putative flexible waterrrows: coupled GluL212 protonation, QB reduction, and QA oxidation in 1.1 ms. (B)

events. In particular, it would be experimentally challenging to cal-culate the 4C O vibration of QB in this scenario and to calculate andmeasure the continuum bands, which would indicate the presenceof a protonated water chain.

Conclusion

In this study, we have shown that a histidine, most likelyHisL190, is successively deprotonated and reprotonated in theintermediate X during the QA

−QB/QAQB− transition. The proton

appears to be shared between HisL190, a water molecule, and QB.This induces protonation of GluL212 via a protonated water chainand creates the electrostatic environment around the QB site nec-essary for the final electron transfer step from QA to QB.

Acknowledgments

This work was supported by the SFB480-C3 grant from theDeutsche Forschungsgemeinschaft. D.O. gratefully acknowledges afellowship from the Alexander von Humboldt Foundation. J.M.S.gratefully acknowledges a fellowship from the Ruhr UniversityResearch School and the Günther and Wilhelm Esser Founda-tion. The authors wish to thank Yvonne Fretter for technicalassistance in protein production and purification. Thanks also toBettina Warscheid and Heike Piechura for performing the massspectrometry.

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 micro-gram 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. Bindingsites of quinones in photosynthetic bacterial reaction centers investigated by

al of C

B

B

C

E

F

G

H

H

H

H

H

H

H

I

Wraight, C.A., 2004. Proton and electron transfer in the acceptor quinone complexof photosynthetic reaction centers from Rhodobacter spaeroides. Front. Biosci. 9,309–337.

D. Onidas et al. / European Journ

light-induced FTIR difference spectroscopy: symmetry of the carbonyl inter-actions and close equivalence of the QB vibrations in Rhodobacter sphaeroidesand Rhodopseudomonas viridis probed by isotope labeling. Biochemistry 34,11606–11616.

reton, J., Richaud, P., Verméglio, A., Nabedryk, E., 2001. Infrared vibrational modesof histidine ligands of the primary electron donor and of QA in purple photo-synthetic bacteria. In: PS2001Proceedings of the 12th International Congress onPhotosynthesis, S7-002. CSIRO Publishing, Collingwood, Victoria, Australia.

rudler, R., de Groot, H.J., van Liemt, W.B., Gast, P., Hoff, A.J., Lugtenburg, J., Gerwert,K., 1995. FTIR spectroscopy shows weak symmetric hydrogen bonding of theQB carbonyl groups in Rhodobacter sphaeroides R26 reaction centres. FEBS Lett.370, 88–92.

hirino, A.J., Lous, E.J., Huber, M., Allen, J.P., Schenck, C.C., Paddock, M.L., Feher, G.,Rees, D.C., 1994. Crystallographic analyses of site-directed mutants of the pho-tosynthetic reaction center from Rhodobacter sphaeroides. Biochemistry 33,4584–4593.

rmler, U., Fritzsch, G., Buchanan, S.K., Michel, H., 1994. Structure of the photo-synthetic reaction centre from Rhodobacter sphaeroides at 2.65 A resolution:cofactors and protein–cofactor interactions. Structure 2 (10), 925–936.

archaus, J.W., Oesterhelt, D., 1989. A Rhodobacter sphaeroides puf L, M and X dele-tion mutant and its complementation in trans with a 5.3 kb puf operon shuttlefragment. EMBO J. 8 (1), 47–54.

raige, M.S., Feher, G., Okamura, M.Y., 1998. Conformational gating of the electrontransfer reaction QA

−QB → QAQB− in bacterial reaction centers of Rhodobacter

sphaeroides determined by a driving force assay. Proc. Natl. Acad. Sci. U.S.A. 95,11679–11684.

asegawa, K., Ono, T.-A., Noguchi, T., 2000. Vibrational spectra and ab initio DFTcalculations of 4-methylimidazole and its different protonation forms: infraredand Raman markers of the protonation state of a histidine side chain. J. Phys.Chem. 104, 4253–4265.

ermes, S., Bremm, O., Garczarek, F., Derrien, V., Liebisch, P., Loja, P., Sebban,P., Gerwert, K., Haumann, M., 2006a. A time-resolved iron-specific X-rayabsorption experiment yields no evidence for an Fe2+ → Fe3+ transition duringQA

− → QB electron transfer in the photosynthetic reaction center. Biochemistry45, 353–359.

ermes, S., Stachnik, J.M., Onidas, D., Remy, A., Hofmann, E., Gerwert, K., 2006b. Pro-ton uptake in the reaction center mutant L210DN from Rhodobacter sphaeroidesvia protonated water molecules. Biochemistry 45, 13741–13749.

essling, B., Souvignier, G., Gerwert, K., 1993. A modelindependent approach toassigning bacteriorhodopsin’s intramolecular reactions to photocycle interme-diates. Biophys. J. 65, 1929–1941.

ienerwadel, R., Berthomieu, C., 1995. Bicarbonate binding to the non-heme ironof photosystem II investigated by Fourier transform infrared difference spec-troscopy and 13C-labeled bicarbonate. Biochemistry 34, 16288–16297.

ienerwadel, R., Grzybek, S., Fogel, C., Kreutz, W., Okamura, M.Y., Paddock, M.L., Bre-ton, J., Nabedryk, E., Mäntele, W., 1995. Protonation of Glu L212 following QB

−

formation in the photosynthetic reaction center of Rhodobacter sphaeroides: evi-

dence from time-resolved infrared spectroscopy. Biochemistry 34, 2832–2843.

olzapfel, W., Finkele, U., Kaiser, W., Oesterhelt, D., Scheer, H., Stilz, H.U., Zinth,W., 1990. Initial electron-transfer in the reaction center from Rhodobactersphaeroides. Proc. Natl. Acad. Sci. U.S.A. 87, 5168–5172.

waki, M., Yakolev, G., Hirst, J., Osyczka, A., Dutton, P.L., Marshall, D., Rich, P.P.,2005. Direct observation of redox-linked histidine protonation changes in the

ell Biology 89 (2010) 983–989 989

iron-sulfur protein of the cytochrome bc1 complex by ATR-FTIR spectroscopy.Biochemistry 44, 4230–4237.

Koepke, J., Krammer, E.-M., Klingen, A.R., Sebban, P., Ullmann, G.M., Fritzsch, G., 2007.pH modulates the quinone position in the photosynthetic reaction center fromRhodobacter sphaeroides in the neutral and charge separated states. J. Mol. Biol.371 (2), 396–409.

Laemmli, U.K., 1970. Cleavage of structural proteins during the assembly of the headof bacteriophage T4. Nature 227 (5259), 680–685.

Li, J., Gilroy, D., Tiede, D.M., Gunner, M.R., 1998. Kinetic phases in the electrontransfer from P+QA

−QB to PQAQB− and the associated processes in Rhodobacter

sphaeroides R-26 reaction centers. Biochemistry 37, 2818–2829.Li, J., Takahashi, E., Gunner, M.R., 2000. −�G0(AB) and pH dependence of the electron

transfer from P+QA−QB to PQAQB

− in Rhodobacter sphaeroides reaction centers.Biochemistry 39, 7445–7454.

Nabedryk, E., Breton, J., Hienerwadel, R., Fogel, C., Mäntele, W., Paddock, M.L.,Okamura, M.Y., 1995. Fourier transform infrared difference spectroscopy ofsecondary quinone acceptor photoreduction in proton transfer mutants ofRhodobacter sphaeroides. Biochemistry 34, 14722–14732.

Okamura, M.Y., Paddock, M.L., Graige, M.S., Feher, G., 2000. Proton and electrontransfer in bacterial reaction centers. Biochim. Biophys. Acta 1458, 148–163.

Paddock, M.L., Feher, G., Okamura, M.Y., 2003. Proton transfer pathways and mech-anism in bacterial reaction centers. FEBS Lett. 555, 45–50.

Raap, J., Winkel, C., de Wit, A.H., van Houten, A.H., Hoff, A.J., Lugtenburg, J., 1990.Mass spectrometric determination of isotopically labeled tyrosines and tryp-tophans in photosynthetic reaction centers of Rhodobacter sphaeroides R-26.Anal. Biochem. 191 (1), 9–15.

Remy, A., 2002. Der QA−QB → QAQB

− Übergang im bakteriellen photosynthetischenReaktionszentrum von Rhodobacter sphaeroides (The QA

−QB → QAQB− transition

in the bacterial photosynthetic reaction centre of Rhodobacter sphaeroides). The-sis of Ruhr-University Bochum, Bochum, Germany.

Remy, A., Gerwert, K., 2003. Coupling of light-induced electron transfer to protonuptake in photosynthesis. Nat. Struct. Biol. 10, 637–644.

Stowell, M.H.B., McPhillips, T.M., Rees, D.C., Soltis, S.M., Abresch, E., Feher, G., 1997.Light induced structural changes in photosynthetic reaction center: Implicationsfor mechanism of electron–proton transfer. Science 276, 812–816.

Tiede, D.M., Vazquez, J., Cordova, J., Marone, P.A., 1996. Time-resolved elec-trochromism associated with the formation of quinone anions in theRhodobacter sphaeroides R26 reaction center. Biochemistry 35, 10763–10775.

Warscheid, B., Brucker, S., Kallenbach, A., Meyer, H.E., Gerwert, K., Kötting, C., 2008.Systematic approach to group-specific isotopic labeling of proteins for vibra-tional spectroscopy. Vib. Spectrosc. 48, 28–36.

Wolpert, M., Hellwig, P., 2006. Infrared spectra and molar absorption coefficients ofthe 20 alpha amino acids in aqueous solutions in the spectral range from 1800to 500 cm(−1). Spectrochim. Acta A Mol. Biomol. Spectrosc. 64 (4), 987–1001.

Zouni, A., Witt, H.T., Kern, J., Fromme, P., Krauss, N., Saenger, W., Orth, P., 2001. Crystalstructure of photosystem II from Synechococcus elongatus at 3.8 A resolution.Nature 409, 739–743.