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. 2003 Mar;84(3):2090–2098. doi: 10.1016/S0006-3495(03)75016-4

The Position of QB in the Photosynthetic Reaction Center Depends on pH: A Theoretical Analysis of the Proton Uptake upon QB Reduction

Antoine Taly *,†, Pierre Sebban †, Jeremy C Smith *, G Matthias Ullmann *
PMCID: PMC1302777  PMID: 12609910

Abstract

Electrostatics-based calculations have been performed to examine the proton uptake upon reduction of the terminal electron acceptor QB in the photosynthetic reaction center of Rhodobacter sphaeroides as a function of pH and the associated conformational equilibrium. Two crystal structures of the reaction center were considered: one structure was determined in the dark and the other under illumination. In the two structures, the QB was found in two different positions, proximal or distal to the nonheme iron. Because QB was found mainly in the distal position in the dark and only in the proximal position under illumination, the two positions have been attributed mostly to the oxidized and the reduced forms of QB, respectively. We calculated the proton uptake upon QB reduction by four different models. In the first model, QB is allowed to equilibrate between the two positions with either oxidation state. This equilibrium was allowed to vary with pH. In the other three models the distribution of QB between the proximal position and the distal position was pH-independent, with QB occupying only the distal position or only the proximal position or populating the two positions with a fixed ratio. Only the first model, which includes the pH-dependent conformational equilibrium, reproduces both the experimentally measured pH dependence of the proton uptake and the crystallographically observed conformational equilibrium at pH 8. From this model, we find that QB occupies only the distal position below pH 6.5 and only the proximal position above pH 9.0 in both oxidation states. Between these pH values both positions are partially occupied. The reduced QB has a higher occupancy in the proximal position than the oxidized QB. In summary, the present results indicate that the conformational equilibrium of QB depends not only on the redox state of QB, but also on the pH value of the solution.

INTRODUCTION

The photosynthetic reaction center (RC) is the pigment-protein complex that performs the initial steps of conversion of light energy into electrochemical energy for ATP synthesis (Okamura et al., 2000; Sebban et al., 1995a). The structures of the RCs from Rhodobacter (Rb.) sphaeroides (Chang et al., 1991; Ermler et al., 1994; McAuley et al., 2000; Stowell et al., 1997) and Rhodopseudomonas (Rps.) viridis (Deisenhofer et al., 1985; Deisenhofer and Michel 1989; Lancaster et al., 2000) have been determined up to a resolution of 2.1 and 2.0 Å, respectively.

The bacterial RC of Rb. sphaeroides is composed of three subunits: L, M and H. The L and M subunits have pseudotwofold symmetry. Both the L and M subunits consist of five transmembrane helices. The H subunit caps the RC on the cytoplasmic side and possesses a single N-terminal transmembrane helix. The RC binds several cofactors: a bacteriochlorophyll dimer, two monomeric bacteriochlorophylls, two bacteriopheophytins, two quinones (QA and QB), a nonheme iron, and a carotenoid. The nonheme iron lies between the two quinone molecules. The primary electron donor, a bacteriochlorophyll dimer called the special pair, is located near the periplasmic surface of the complex, and the terminal electron acceptor, a quinone called QB, is located near the cytoplasmic side.

Electron transfer from the special pair to QB is initiated by the absorption of light, which induces the excitation of the special pair to its lowest excited electronic state. The electron is subsequently transferred in 200 ps to QA via a monomeric chlorophyll and a pheophytin. QA− is oxidized in 20–200 μs by electron transfer to QB (Li et al., 1998; Tiede et al., 1996). In the RCs of Rb. sphaeroides, QA and QB are both ubiquinone molecules. However, these two ubiquinone molecules have different properties and different functions. The QB binding pocket is richer in polar and ionizable residues than that of QA. Although QA is a one-electron acceptor and does not protonate directly, QB accepts two electrons and two protons to form the reduced QBH2 molecule. QB is bound at the level of the lipid headgroups at the cytoplasmic side of the membrane and has no direct contact with the aqueous environment. Protons are delivered from the cytoplasm to QB by one or more pathways composed of interdependent hydrogen-bond networks involving titratable residues and water molecules (Baciou and Michel, 1995; Ermler et al., 1994; Gerencser et al., 2002; Lancaster and Michel, 1997; Lancaster et al., 1996; Miksovska et al., 1997; Paddock et al., 2001).

The first reductions of QA and of QB are accompanied by pKa shifts of residues that interact with the semiquinone species (Wraight, 1979). The reductions induce substoichiometric proton uptake by the protein. The number of protons taken up by the protein upon reduction of the quinones is an observable directly dependent on the energetics of the system and is also intimately coupled to the thermodynamics of the QA−→QB electron transfer process (Okamura et al., 2000; Onufriev et al., 2001). Moreover, it has been proposed that proton uptake and rearrangements after QA− formation could be dynamically coupled to the interquinone electron transfer reaction and may gate this reaction (Brzezinski et al., 1992; Maróti and Osváth, 1997; Tiede and Hanson, 1992). The pH dependence of the proton uptake associated with the formation of QA− and QB− in wild type RCs have been determined for Rb. sphaeroides (Maroti and Wraight, 1988; McPherson et al., 1988; Tandori et al., 2002) and Rb. capsulatus (Sebban et al., 1995b).

Using x-ray structural analysis, it has been shown that a major conformational difference exists between the RC handled in the dark (the ground state) or under illumination (the charge-separated state) (Stowell et al., 1997). The main difference between the two structures concerns QB itself, which was found in two different positions ∼4.5 Å apart. In the dark-adapted state in which QB is oxidized, QB is found mainly in the distal position and only a small percentage in the proximal position. Under illumination, i.e., when QB is reduced, QB is seen only in the proximal position. The crystal was grown at pH = 8 (Allen, 1994). The reaction center structures with proximal or distal QB are called RCprox and RCdist, respectively (Lancaster and Michel, 1997). A similar conformational equilibrium was found for the RC of Rps. viridis (Lancaster, 1999a). A schematic representation of this crystallographically observed equilibrium is shown in Fig. 1.

FIGURE 1.

FIGURE 1

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Illustration of the crystallographically determined equilibrium (Stowell et al., 1997). The left side represents the structure of dark-adapted RCs, in which QB is oxidized. QB is seen in two positions: proximal and distal. The structures with proximal and distal QB are called RCprox and RCdist, respectively. The right side represents the structure for light-exposed RCs, in which QB is reduced. No electron density is observed experimentally for RCdist.

Conformational changes can shift pKa values of residues in proteins (Beroza and Case, 1998; Gunner and Alexov, 2000; Huang et al., 2002; Mulkidjanian, 1999; Rabenstein and Knapp, 2001). It is therefore interesting to investigate the effect of pH and thus protonation state changes on the conformational equilibria associated with pKa switching. Conformational equilibria play a central role in the physiological function of many proteins (Graige et al., 1998; Huang et al., 2002; Mulkidjanian, 1999; Rabenstein and Knapp, 2001). For the RC, it was, for instance, proposed that a conformational equilibrium participates in the gating of the first electron transfer between QA and QB (Graige et al., 1998). A theoretical understanding of the pH dependence of conformation equilibria is therefore of general interest in protein biophysics. This understanding can be approached with the use of the Poisson-Boltzmann equation in which the protein atoms are explicitly represented by partial charges and the environmental effect are included by a continuum description (Honig and Nicholls, 1995; Honig et al., 1989; Sharp et al., 1995; Sharp and Honig, 1990; Yang et al., 1993). This approach allows the computation of the electrostatic potential at any point inside and outside the protein. The electrostatic potential depends on and determines the protonation of the individual residues. Combining Poisson-Boltzmann calculations with Monte Carlo sampling of protonation states allows calculating the overall proton uptake to be performed, together with a decomposition of the contributing residues.

In the calculations presented here, we investigate how the proton uptake upon QB reduction in the Rb. sphaeroides RC depends on the pH and on the conformational equilibrium of QB found experimentally (Stowell et al., 1997). We show that a model, in which the equilibrium between the conformations RCprox and RCdist varies with pH, reproduces the experimentally measured pH dependence of the proton uptake (Tandori et al., 2002) as well as the occupation of RCprox and RCdist observed in the crystallographic study at pH = 8 (Stowell et al., 1997). In the model, the populations of the two conformations in the ground and charge-separated states of the RC are pH dependent. The results of the study provide insight into the balance between the global protein electrostatics and conformational equilibrium of a protein, and how conformational equilibria are controlled by pH.

METHODS

Structure preparation

X-ray structures used

The x-ray structures of the dark-adapted (PDB entry 1AIJ) and light-exposed (PDB entry 1AIG) RCs from Rb. sphaeroides with resolutions of 2.2 and 2.6 Å (Stowell et al., 1997), respectively, were used in this study. 1AIG was used for all states in which QB is in the proximal position; 1AIJ was used for all states in which QB is in the distal position.

Structure preparation

Two RCs are present in the asymmetric unit of the crystal. We consider only the first RC structure of the PDB entries 1AIJ and 1AIG, not the second. The second RC in the asymmetric unit is less complete than the first. All explicit water and detergent molecules were removed. The influence of water was represented using a dielectric constant of 80 (Adcock et al., 1998; Baptista and Soares, 2001; Gibas and Subramaniam, 1996; Lancaster et al., 1996; Rabenstein et al., 1998; Teixeira et al., 2002). The use of lower dielectric constants (e.g. 30) inside cavities does not influence protonation probability calculations significantly (Adcock et al., 1998). Therefore, we did not consider this effect. Most of the nonpolar hydrogen atoms were considered as one atom together with the heavy atoms to which they are bound (the extended atom representation). For the quinones, the bacteriochlorophylls, and the bacteriopheophytins all hydrogens were treated explicitly. Polar hydrogens, i.e., those bound to oxygen, nitrogen, or sulfur atoms, were also treated explicitly, except for the acidic hydrogens of protonated carboxylate groups which were represented by symmetrical charge adjustment of the two carboxyl oxygen atoms (Rabenstein et al., 1998). Coordinates of explicitly treated hydrogen atoms were generated with the HBUILD module (Brunger and Karplus, 1988) in CHARMM (Brooks et al., 1983). The hydrogen atom positions were energy optimized with the heavy-atom positions fixed. For this optimization, all titratable groups were in their standard protonation states (i.e., the aspartate, glutamate and the C-termini were unprotonated; the arginine, cysteine, histidine, lysine, tyrosine and the N-termini were protonated), and both quinones were in their oxidized (uncharged) state. The hydrogen atom positions were kept fixed during the electrostatic calculations. The continuum representation of the water and symmetrical distribution of the charges over the protonatable groups mimics the mobility of the hydrogen atoms and the water molecules well (Gibas and Subramaniam, 1996). This representation is computationally much less demanding than treating these effects explicitly. We used the same atomic partial charges as in previous calculations (Rabenstein et al., 1998, 2000). The charges of bacteriopheophytin and bacteriochlorphyll, which have not been published before, are listed in Supplementary Material.

Proton uptake calculation

Calculation of protonation probabilities

Each protonation state of a protein can be characterized by a protonation state vector Inline graphic, where the components Inline graphic are 1 or 0 depending on whether group μ is protonated or not. The superscripts n and k designate the protonation state and the conformation of the protein, respectively. The energy Gn,k of a protonation state n of the protein in a conformation k is given by Eq. 1 (Bashford and Karplus, 1990; Ullmann and Knapp, 1999).

graphic file with name M3.gif (1)

where Inline graphic is the unitless formal charge of the deprotonated form of group μ, i.e., −1 for acids and 0 for bases, and Inline graphic is the reference protonation state of group μ; Inline graphic is the experimentally known pKa value of a model compound of the titratable group (N-formyl N-methylamide derivatives of the respective amino acids) in aqueous solution (Tanford and Roxby, 1972); Inline graphic is the shift of the model compound pKa value of the titratable groups due to the different solvation environment inside the protein (changed dielectric environment and interaction with non-titrating charges); Inline graphic is the electrostatic interaction between the titratable groups μ and ν in the conformation k if both are charged; δk is 1 or 0 depending on whether the protein is in conformation k or not; R is the universal gas constant, and T is the temperature. Inline graphic is the free energy difference between conformation k and the reference conformation k = 0 in which all sites are in the reference protonation state (Eq. 2). In the present case, the reference conformation k = 0 is that with a proximal quinone.

graphic file with name M10.gif (2)

This energy difference refers to the energy of the specific protein conformation and for a specific redox state of QB. Here, ΔGconf has two different values, one for each redox state of QB: Inline graphic when QB is reduced and Inline graphic when QB is oxidized. As discussed by others (Rabenstein and Knapp, 2001), the value of ΔGconf is composed of different contributions, such as, for example, van der Waals interactions, and Coulombic interactions between nontitratable groups and titratable groups (the latter being only considered in the reference protonation state), and torsion energies. Accurate determination using theory of each of these contributions, and thus ΔGconf, is difficult. Therefore, it is common practice to treat ΔGconf as an adjustable parameter to reproduce experimental data. Here ΔGconf was kept constant with pH.

The terms Inline graphic and Inline graphic were calculated from the linearized Poisson-Boltzmann equation of a molecular system using a finite difference method with the program MEAD (Bashford and Gerwert, 1992). The Poisson-Boltzmann equation was solved using a three-step grid-focusing procedure (Bashford and Gerwert, 1992; Klapper et al., 1986; Rabenstein et al., 1998) with an initial 250-Å cube with a 2.5-Å lattice spacing centered at the protein, followed by 100-Å cube with a 1.0-Å lattice spacing, and a 45-Å cube with 0.3-Å lattice spacing, both centered at the titratable group. We used an ionic strength of 100 mM, an ion exclusion layer of 2 Å, and a solvent probe radius of 1.4 Å. The dielectric constant of the protein was set to ɛP = 4 and the dielectric constant of the solvent (outside the protein and within protein cavities) was set to ɛS = 80.

The average protonation probability of each titratable group was calculated by a Monte Carlo procedure (Beroza et al., 1991) using the program Karlsberg (Rabenstein and Knapp, 2001; Rabenstein et al., 2000). For the histidines, two tautomers were considered explicitly. All other titratable groups were treated by a single tautomer, which represented an average over all possible tautomers. In previous studies, this approach gave good agreement with experimentally determined pKa values (Bashford et al., 1993; Rabenstein and Knapp, 2001; Rabenstein et al., 1998, 2000; Ullmann, 2000). The Monte Carlo sampling was sufficient to reach a standard deviation of less than 0.01 proton at each individual titratable group. Most of the standard deviations were much smaller than 0.01. The sum of the standard deviations of all protonation probabilities was ∼0.02 proton.

Proton uptake calculation

The protonation probabilities of the 172 titratable residues were computed for the states QAQB and QAQB−. The protonation probability difference between the states QAQB and QAQB− was directly compared to the corresponding experimental data. The experimental pH dependence of the proton uptake determined by Tandori et al., 2002 and McPherson et al., 1988 are very similar. In plots presented in this paper, the experimental data from Tandori et al., 2002 are used for comparison with the calculations. Four different models were used for the proton uptake calculations:

In Model 1 (Fig. 2 a), the conformational equilibrium and redox states are pH dependent. Four possible redox and conformational states of the RC are included: oxidized QB in the proximal position (Inline graphic), reduced QB in the proximal position (Inline graphic), oxidized QB in the distal position (Inline graphic), and reduced QB in the distal position (Inline graphic). In the model, each of these redox and conformational states exists in 2N protonation states where N is the number of protonation sites. These states are in thermodynamic equilibrium, i.e., they are populated with the Monte Carlo method according to Boltzmann statistics. ΔGconf was adjusted such that both the experimentally determined pH dependence of the proton uptake (Tandori et al., 2002) and the crystallographically determined equilibrium between the two structures observed at pH = 8 (Stowell et al., 1997) were simultaneously reproduced. This agreement was achieved as follows: the difference between the calculated and experimental pH dependence of the proton uptake was minimized subject to two constraints: i), the occupancy of the RCprox conformation was constrained to be lower than 50% when QB is oxidized, and ii), when QB is reduced, the occupancy of the RCprox structure should be at least 70%. These two constraints ensure that the results are consistent with the observations made crystallographically (Stowell et al., 1997). The values obtained for the conformational energy difference are Inline graphic = 1.20 eV and Inline graphic = 1.23 eV. The values found here are of the same order as in previous studies (Rabenstein and Knapp, 2001).

FIGURE 2.

FIGURE 2

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Illustration of the four models used to test the relationship between RC conformations and proton uptake upon reduction of QB. (a) Model 1 describes a pH-dependent conformational equilibrium between RCprox and RCdist for QB and QB−. (b) Model 2 uses the populations of RCprox and RCdist for QB and QB− found with the first model at pH 8 and keeps the ratio of the different conformations constant over the whole pH range. (c) Model 3 uses only the RCprox structure for both redox states of QB over the whole pH range. (d) Model 4 uses only the RCdist structure for both redox states of QB over the whole pH range.

Model 2 (Fig. 2 b) allows one to test if the conformational equilibrium is pH dependent or not. To do this, the populations of RCprox and RCdist are fixed and constant over the whole pH range for each quinone redox state. The populations used are those determined at pH = 8 with Model 1 (i.e., those consistent with the crystallographic results obtained at pH 8).

In Model 3 (Fig. 2 c) and Model 4 (Fig. 2 d) only a single structure, RCprox or RCdist respectively, is used in the calculations. Therefore, in both of these models QB is in the same position over the whole pH range, whether reduced or not. Model 3 and Model 4 are used to test if a single conformation is sufficient to reproduce the experimental pH dependence of the proton uptake.

RESULTS AND DISCUSSION

The aim of the present work is to understand the pH dependence of the proton uptake associated with the reduction of QB and its relation to the experimentally observed conformational equilibrium. Four models were compared to two sets of experimental data: the conformational equilibrium between the RCprox and RCdist structures found at pH = 8 by x-ray crystallography (Stowell et al., 1997) and the pH dependence of the proton uptake upon QB reduction (Tandori et al., 2002).

The structural equilibrium found crystallographically (Stowell et al., 1997) is schematically shown in Fig. 1. In the dark-adapted RC (left side of Fig. 1), QB is found in the two positions with a majority in the distal position. In contrast, no electron density was observed for QB in the distal position in the light-exposed RC. This observation implies that under light illumination when QB is reduced, the proportion of RC with distal QB is very low at pH 8 where the structure was determined.

Proton uptake calculations with different models

The proton uptake upon QB reduction was calculated with four different models as described in the Methods section. The results are compared with the experimentally measured proton uptake curves.

Model 1: pH- and redox-dependent equilibrium between RCprox and RCdist

Model 1 is shown in Fig. 2 a. In this model, the RC adapts two conformations, RCprox and RCdist, in both oxidation states of QB. The equilibrium between the two conformations was adjusted to fit the experimental proton uptake data by varying ΔGconf. The model implies that when QB is neutral both structures are equally populated at pH = 8. In contrast, RCprox is 70% occupied at pH = 8 when QB is reduced. Changing the population probabilities of the two positions to other rations which are also in agreement with crystallographic data led to the same behavior of the proton uptake curve, i.e., first a decrease of the proton uptake followed by an increase, but with worse overall agreement with the proton uptake data. The equilibrium found from the fits describes the pH dependence of the proton uptake and the crystallographically observed conformational equilibrium well.

In the neutral pH range, the population of RCprox is higher when QB is reduced than when it is oxidized, as was imposed for consistency with the x-ray observations (Stowell et al., 1997). The shift of the equilibrium between RCprox and RCdist upon QB reduction is somewhat smaller in the calculations than seen crystallographically. However, the difference between the calculated equilibrium and the one seen by x-ray crystallography corresponds to a small energy difference of the order of the thermal fluctuation energy (kBT ∼ 0.6 kcal/mol), which is well within the error of the method.

The experimental pH dependence of the proton uptake presented in Fig. 3 a decreases from pH 6 to 8 and increases above pH 8 to reach a plateau above pH 9. The calculated proton uptake reproduces this shape. Therefore, Model 1, which allows a pH-dependent structural equilibrium between RCprox and RCdist for QB and QB−, is capable of satisfactorily describing the pH dependence of the proton uptake.

FIGURE 3.

FIGURE 3

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(a) Proton uptake upon reduction of QB, calculated using Model 1, which considers a conformational equilibrium between RCprox and RCdist for QB and QB− is shown by the black line. The experimental proton uptake at the different pH values are shown by gray circles. (b) Equilibrium between RCprox and RCdist structures. The calculated ration RCprox/(RCprox + RCdist) is shown for oxidized (dashed line) and reduced (solid line) QB.

Fig. 3 b shows the pH dependence of the equilibrium between RCprox and RCdist as a function of pH for both redox states of QB, resulting from Model 1. In both states, QAQB and QAQB−, the occupancy of the RCprox structure increases with increasing pH.

The results indicate that the structural equilibrium between the conformations RCprox and RCdist depends on both the redox state of QB and the pH value of the solution. Model 1 indicates that the structural transition is controlled by the redox state of QB only in the pH range between 6.5 and 8.5. At lower or higher pH the equilibrium is totally shifted to the conformations RCdist or RCprox, respectively.

It is known from experiments that QB is loosely bound at high pH. The experimental proton uptake data, with which we compare our results, are corrected for this effect (Tandori et al., 2002). Our calculations indicate that at high pH the proximal position of QB is favored relative to the distal position independent of the redox state of QB and the occupation of the QB site.

Decomposition of the proton uptake in Model 1

The global proton uptake can be decomposed into two major contributions, which are shown in Fig. 4. The proton uptake is mainly due to residues Glu-L212 and Asp-L213. At pH 7 the proton uptake due to Glu-L212 is calculated to be 0.54, in reasonable agreement with the value of ∼0.3–0.4 obtained from FTIR experiments (Nabedryk et al., 1995). According to our results, the residues Asp-L210 and Glu-H173 do not change their protonation probability significantly upon QB reduction. In the neutral pH range, the difference between the proton uptake of the residues Glu-L212 and Asp-L213 and the total proton uptake arises from the conformational change that is accompanied by small changes of protonation probabilities of several residues (Arg-M136, Asp-M17, Glu-H33, Glu-M236, His-H116, His-H118, Lys-H50, Lys-H52, Lys-H136, Lys-H187, Lys-H222). The residues that are in contact with the membrane region do not participate in the proton uptake.

FIGURE 4.

FIGURE 4

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Decomposition of the proton uptake obtained with Model 1. Experimental proton uptake is shown by gray circles, total proton uptake by a solid line, contribution of residues GLU-L212 and ASP-L213 by a dashed line, and the contribution of the remaining residues, which are listed in the main text, are shown by a dotted line.

The total protonation probability of the RC is higher in conformation RCdist than in the conformation RCprox. Thus, RCdist is stabilized at low pH and RCprox at high pH. However, Glu-L212 and Asp-L213 are more protonated in the RCprox conformation (Table 1) even at low pH. Other residues listed in the previous paragraph change their protonation because of the conformational transition and thus compensate for this effect. The titration of Glu-L212 and Asp-L213 is irregular (Table 1). Such an irregular titration behavior is indicative of complicated electrostatic interaction and has been also observed for other molecules (Onufriev et al., 2001; Sudmeier and Reilley, 1964; Zuiderweg et al., 1979).

TABLE 1.

Protonation probability of Glu-L212 and Asp-L213 when QB is reduced calculated for the conformations RCdist and RCprox and for Model 1

pH
Residue QB position 6.0 8.0 10.0
Glu-L212 RCprox 1.0 1.0 1.0
RCdist 0.78 0.3 0.27
Model 1 0.78 0.82 1.0
Asp-L213 RCprox 1.0 0.99 0.92
RCdist 0.89 0.85 0.77
Model 1 0.89 0.94 0.92
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Model 2: pH-independent, redox-driven transition between RCprox and RCdist

This model was designed to test the hypothesis that structural rearrangement might be driven solely by the change of redox state of QB. According to this hypothesis, the position of QB changes when QB is reduced, independent of the pH. The calculations were therefore performed with the populations of RCprox and RCdist imposed at the values found with Model 1 at pH = 8 over the whole pH range, i.e., 50% RCprox when QB is oxidized and 70% RCprox when QB is reduced. The proton uptake in the pH range from 6 to 10 calculated using Model 2 is shown in Fig. 5 a. Clearly, this model cannot qualitatively reproduce the variation with pH of the experimental proton uptake upon QB reduction. The calculated proton uptake is in good agreement with experiments in the pH range from 6.5 to 8.5. However, outside the range from 6.5 to 8.5, there is clear deviation from experiment. This disagreement is a further indication that the conformational equilibrium between RCprox and RCdist is pH dependent, and that QB changes its equilibrium position after reduction only in the pH range from 6.5 to 8.5.

FIGURE 5.

FIGURE 5

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Proton uptake of the RC upon reduction of QB. Comparison of calculated (solid line) and experimental data (circles). The calculations were performed with (a) Model 2. (b) Model 3. (c) Model 4. The proton uptake of Model 1 is shown as a dotted line for comparison.

Model 3: fixed conformation RCprox

In this model, the RCprox structure was used for both redox states of QB over the whole pH range (Fig. 2 c). The pH dependence of the proton uptake upon reduction of QB calculated using this model is displayed in Fig. 5 b. This model is in agreement with experiments only around pH 6 and between pH 9 and 10 but cannot reproduce either the experimentally observed decrease in proton uptake between pH 6 and 8 or the increase between pH 8 and 9. The agreement between calculated and experimental proton uptake in the pH range from pH 9 to 10 is consistent with the results obtained from Model 1, because the RCprox structure satisfactorily represents the RC in both redox states in the pH range from 9 to 10. According to Model 1, only RCprox is populated above pH 9.

Model 4: fixed conformation RCdist

In Model 4, the RCdist structure was used for both redox states of QB over the whole pH range (Fig. 2 d). The pH-dependent proton uptake calculated from this model is shown in Fig. 5 c. The model reproduces well the experimental proton uptake below pH = 8. However, above pH = 8, the results from Model 4 do not even qualitatively follow the experimental data. This finding is again in agreement with Model 1, because the RC populates only the RCdist conformation at low pH for both QB and QB−.

Comparison of the four models

Only Model 1, which includes a pH-dependent structural equilibrium between the RCprox and RCdist, describes the proton uptake experiments satisfactorily over the whole pH range. We therefore conclude that a pH-dependent structural equilibrium between RCprox and RCdist is necessary to describe the pH dependence of the proton uptake upon QB reduction. Models 2, 3, and 4 describe the experimental data well over limited pH ranges over which they are found to be valid approximations to Model 1.

Relation to previous calculations

Several theoretical studies on protonation probabilities and conformational changes in the QB pocket of different RCs have be done before (Alexov et al., 2000; Alexov and Gunner, 1999; Grafton and Wheeler, 1999; Lancaster et al., 1996; Lancaster, 1999b; Rabenstein et al., 1998, 2000; Walden and Wheeler, 2002; Zachariae and Lancaster, 2001). However, none of these studies considered the pH dependence of the conformational transition between RCprox and RCdist. Here, we make that connection and corroborate our calculation by reproducing experimental proton uptake measurements.

A previous study used only one structure in the evaluation of the proton uptake (Beroza et al., 1995). In this structure (PDB entry 4RCR), QB is in the proximal site. The proton uptake calculated in this study (Beroza et al., 1995) is within 0.05 proton of the results obtained in the present work using only the RCprox structure over the whole pH range (i.e., Model 3).

The way flexibility is treated in Model 1 differs from previous calculations in which the flexibility was introduced by allowing the side chains of 26 residues to occupy the different conformations found in the different x-ray structures of the RC protein (Alexov and Gunner, 1999). In addition, QB was allowed to occupy the distal and proximal positions. This model involves a large number of possible conformational substates and thus also many parameters to describe them. The calculations done with this model reproduced the pH dependence of proton uptake, but the pH dependence of the quinone position occupancy was not reported.

Interestingly, according to our calculations, residues Glu-L212 and Asp-L213 are protonated from pH 6 to 10 when QB is reduced and proximal. A study using molecular dynamics simulation has shown that the proximal position of QB− is more stable when both residues Glu-L212 and Asp-L213 are protonated (Grafton and Wheeler, 1999). The present study is therefore in agreement with this work. However, the results obtained in the present study suggest that the position of QB does not depend only on the protonation state of L212 and L213, but also on the protonation state of other residues that trigger the conformational transition between RCprox and RCdist. This finding is also supported by a more recent study (Walden and Wheeler, 2002). It should, however, be mentioned that in those theoretical studies (Grafton and Wheeler, 1999; Walden and Wheeler, 2002) QB occupies the proximal position in an orientation that has never been found crystallographically (McAuley et al., 2000; Zachariae and Lancaster, 2001)

Molecular dynamics simulations of the RC of Rps. viridis have provided evidence supporting the movement of QB between the distal site and the proximal site (Zachariae and Lancaster, 2001). This work showed that the equilibrium between the two binding sites is not only displaced by the reduction of QB to the semiquinone, but also by the preceding reduction of the primary quinone QA and accompanying protonation changes in the protein. The present model supports this idea, because the position of QB is influenced by the protonation states of the residues surrounding QB which may in turn be influenced by the redox state of QA (Zachariae and Lancaster, 2001).

CONCLUSIONS

The aim of the present study was to understand the pH dependence of the proton uptake associated with the reduction of QB. Two experimentally observed conformations of the RC were considered: with QB bound in the proximal or the distal binding site. Comparing the calculated and experimental pH dependence of the proton uptake reveals that a pH-dependent conformational transition is required to reproduce the experimental proton uptake curve. Neither the individual conformations nor a static mixture of the two conformations with a pH-independent population are capable to reproduce the experimental proton uptake profile. The present study presents a new picture in which the position of QB depends not only on the redox state of QB, but also on pH. This hypothesis could be tested experimentally, for instance by x-ray crystallography at different pH values.

The kinetics of the first electron transfer reaction between QA and QB is biphasic (∼20–60 μs and 150–400 μs) (Li et al., 1998; Tiede et al., 1996). Both electron transfer rates are gated, i.e., not limited by the electron transfer process itself but by other processes (Graige et al., 1998; Hoffman and Ratner, 1987; Ullmann et al., 1997; Zhou and Kostic, 1993). The conformational transition of QB from the distal to the proximal sites has been proposed to be one of the rate limiting steps of the first electron transfer (Graige et al., 1998).

Because of the observation that QB occupies the proximal position when it is reduced (Stowell et al., 1997), i.e., after the electron transfer, the proximal position has been suggested to be active for electron transfer and the distal position to be inactive (Graige et al., 1998). However, a proximal position of QB is not necessarily associated with a nongated electron transfer (Ädelroth et al., 2000; Kuglstatter et al., 2001; Tandori et al., 2002). Consequently, when QB is proximal other processes may also gate the first electron transfer from QA to QB. However, if the movement of QB is one of the rate limiting steps, our results imply that the proportion of RCs for which the first electron transfer between QA and QB is gated by the movement of QB will decrease with increasing pH. This idea will be tested in future theoretical and experimental studies.

SUPPLEMENTARY MATERIAL

S1.

Partial charges of Bacteriopheophytin, calculated by a semi-empirical method. Atom numbers are according to the PDB file IAIG

Atom name Charge Atom name Charge Atom name Charge Atom name Charge
NA −0.49 1HAA 0.02 C5 0.00 C3C −0.22
NB −0.5 2HAA 0.02 H8 0.00 CAC −0.11
HNB 0.36 C3A −0.22 H9 0.00 2HAC 0.07
NC −0.35 H3A 0.07 C6 0.00 CBC 0.08
ND 0.52 CMA 0.00 H10 0.00 2HBC 0.00
HND −0.01 1HMA 0.01 H11 0.00 3HBC 0.00
C1A −0.02 2HMA 0.01 C2B −0.04 1HBC 0.00
CHA 0.11 3HMA 0.01 CMB −0.02 C2D 0.05
C4D −0.15 CBA −0.3 1HMB 0.03 CMD 0.05
C1B 0.44 2HBA 0.1 2HMB 0.03 1HMD 0.01
CHB −0.65 1HBA 0.1 3HMB 0.03 2HMD 0.01
HHB 0.21 CGA 0.79 C3B −0.39 3HMD 0.01
C4A 0.56 O1A −0.54 CAB 0.79 C3D −0.43
C1C 0.28 O2A −0.43 OBB −0.52 CAD 0.89
CHC −0.46 C1 0.19 CBB −0.41 OBD −0.55
HHC 0.17 H1 0.01 HB1 0.11 CBD −0.8
C4B 0.4 H2 0.01 HB2 0.11 1HBD 0.3
C1D −0.25 C2 0.00 HB3 0.11 CGD 0.98
CHD −0.2 H3 0.00 C2C 0.17 OID −0.53
HHD 0.2 C3 0.00 H2C 0.02 O2D −0.49
C4C 0.17 C4 0.00 CMC −0.09 CED 0.27
C2A 0.14 H5 0.00 1HMC 0.03 1HED −0.02
H2A 0.06 H6 0.00 2HMC 0.03 2HED −0.02
CAA 0.06 H7 0.00 3HMC 0.03 3HED −0.02
NA 0.11 CAA −0.13 H7 0.00 2HAC 0.08
NB 0.12 1HAA 0.09 C5 0.00 CBC 0.11
NC 0.01 2HAA 0.09 H8 0.00 2HBC 0.00
ND 0.01 C3A −0.04 H9 0.00 3HBC 0.00
C1A 0.03 H3A 0.06 C2B −0.16 1HBC 0.00
CHA −0.22 CMA 0.01 CMB 0.11 C2D 0.18
C4D 0.21 1HMA 0.01 1HMB −0.01 CMD −0.03
C1B −0.05 2HMA 0.01 2HMB −0.01 1HMD 0.03
CHB −0.2 3HMA 0.01 3HMB −0.01 2HMD 0.03
HHB 0.19 CBA −0.21 C3B −0.32 3HMD 0.03
C4A −0.1 2HBA 0.08 CAB 0.65 C3D −0.41
C1C −0.19 1HBA 0.08 OBB −0.57 CAD 0.81
CHC −0.17 CGA 0.77 CBB −0.14 OBD −0.52
HHC 0.2 O1A −0.53 HB1 0.04 CBD −0.73
C4B −0.02 O2A −0.42 HB2 0.04 1HBD 0.29
C1D −0.06 C1 0.15 HB3 0.04 CGD 1.04
CHD −0.13 H1 0.01 C2C 0.24 O1D −0.57
HHD 0.18 H2 0.01 H2C −0.01 O2D −0.5
C4C 0.05 C2 0.00 CMC −0.05 CED 0.19
C2A 0.1 H3 0.0 1HMC 0.02 1HED 0.00
H2A 0.05 C3 0.0 2HMC 0.02 2HED 0.00
CAA −0.13 C4 0.0 3HMC 0.02 3HED 0.00
1HAA 0.09 H5 0.0 C3C −0.12 MG 0.0
2HAA 0.09 H6 0.0 CAC −0.18
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Acknowledgments

We are grateful to Prof. Donald Bashford and to Dr. Bjoern Rabenstein for providing their programs MEAD and Karlsberg, respectively. We thank Dr. Laura Baciou for stimulating discussions.

A.T. is supported by a fellowship from the Deutscher Akademischer Austauschdienst (A/00/00063). G.M.U. is supported by a Emmy-Noether grant of the Deutsche Forschungsgemeinschaft (UL174/2-1). This work was supported by a PROCOPE program (DAAD/CNRS).

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