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. 2022 Nov 14;126(46):9549–9558. doi: 10.1021/acs.jpcb.2c05713

Electron Transfer Route between Quinones in Type-II Reaction Centers

Yu Sugo †, Hiroyuki Tamura †,‡, Hiroshi Ishikita †,‡,*
PMCID: PMC9707520  PMID: 36374126

Abstract

graphic file with name jp2c05713_0005.jpg

In photosynthetic reaction centers from purple bacteria (PbRCs) and photosystem II (PSII), the photoinduced charge separation is terminated by an electron transfer between the primary (QA) and secondary (QB) quinones. Here, we investigate the electron transfer route, calculating the superexchange coupling (HQA–QB) for electron transfer from QA to QB in the protein environment. HQA–QB is significantly larger in PbRC than in PSII. In superexchange electron tunneling, the electron transfer via unoccupied molecular orbitals of the nonheme Fe complex (QA → Fe → QB) is pronounced in PbRC, whereas the electron transfer via occupied molecular orbitals (Fe → QB followed by QA → Fe) is pronounced in PSII. The significantly large HQA–QB is caused by a water molecule that donates the H-bond to the ligand Glu-M234 in PbRC. The corresponding water molecule is absent in PSII due to the existence of D1-Tyr246. HQA–QB increases in response to the Ser-L223···QB H-bond formation caused by an extension of the H-bond network, which facilitates charge delocalization over the QB site. This explains the observed discrepancy in the QA-to-QB electron transfer between PbRC and PSII, despite their structural similarity.

Introduction

Photosynthetic reaction centers from purple bacteria (PbRCs) and photosystem II (PSII), classified as type-II reaction centers, share structural similarities1 and are believed to have evolved from the same ancestor.2,3 The protein subunits L and M form a heterodimeric reaction center in PbRC. Photoinduced charge separation predominantly occurs via bacteriochlorophyll and bacteriopheophytin in subunit L.4,5 The subsequent electron transfer occurs from the primary quinone (QA) of subunit M to the secondary quinone (QB) of subunit L,6 leading to the formation of semiquinone (QB•–) and the proton transfer toward the carbonyl O site of QB via Asp-L213 and Ser-L223 (distal carbonyl site).7−9 The second electron transfer leads to the second reduction of QB followed by the release of the proton from His-L190 toward the carbonyl O site (proximal carbonyl site) of QB, forming QBH2.9 QBH2 releases from the QB binding site, and unprotonated quinone enters the QB binding pocket.10 The first electron transfer occurs with the observed rate constant kAB(1).11 The redox potential (Em) difference between QA and QB is reported to be 30 to 70 mV.6,8,12−14 As kAB(1) is independent of the Em difference between QA and QB, the process includes electron transfer (with the rate constant kET) and rate-limiting “conformational gating”,11 where 1/kET ≪ 1/kAB(1) = 110 μs.11 The conformational gating is absent in the second electron transfer.11 The nonheme Fe complex, composed of one glutamate ligand and four histidine residues, is equidistant from QA and QB.15 It donates H-bonds to QA and QB via His-M219 and His-L190 residues, respectively, thus forming the H-bond network [QA···His-M219···Fe···His-L190···QB]. The depletion or substitution of Fe with other divalent metals does not cause kAB(1) to change significantly.16 The depletion of Fe leads to the protonation of either His-M219 or His-L190 histidine ligands, which partially compensates for the loss of the divalent metal.17 It seems plausible that the nonheme Fe complex is not a prerequisite for rapid QA-to-QB electron transfer in PbRC.16

The cofactor arrangement of PSII resembles that of PbRC. QA and QB are ubiquinone in PbRC from Rhodobacter sphaeroides and plastoquinone in PSII (Figure S1); the low barrier H-bond between His-L190 and QBH– in PbRC9 is also conserved as that between D1-His215 and QBH– in PSII18,19 (note: Rhodobacter sphaeroides was previously known as Rhodopseudomonas sphaeroides and now reclassified as Cereibacter sphaeroides(20)). However, the electron transfer specifics differ for the two type-II reaction centers. The experimentally measured Em difference between QA and QB is ∼200 mV in PSII,21,22 which is more than 3 times that in PbRC. In contrast, the time constant for the first electron transfer from QA to QB is 200–400 μs,23 indicating that electron transfer in PSII is 2–4 times slower than that in PbRC. The QB exchange time in PSII (∼20 ms)24,25 is longer than that in PbRC (∼1 ms),10 partially because the release of QB leads to the transformation of the short helix (D1-Phe260 to D1-Ser264) in PSII,26 which is not conserved in PbRC. The two quinones are bridged via the nonheme Fe complex in PSII; however, the glutamate ligand in PbRC is replaced by the bicarbonate ligand in PSII. The replacement of bicarbonate with formate leads to an inhibition of the electron transfer from QA to QB,27−29 which suggests that the nonheme Fe complex is involved in electron transfer in PSII. The nonheme Fe complex is redox-active in PSII, with Em(Fe2+/Fe3+) = 400 mV,30,31 whereas the redox activity has not been reported for PbRC.

To the best of our knowledge, the discrepancy in electron transfer from QA to QB between the two type-II reaction centers remains unsolved, based on protein structures that have essentially conserved cofactor arrangements and geometries. Here, we investigate the electron transfer route from QA to QB in PbRC and identify the factors that differentiate the mechanism between the two type-II reaction centers using a quantum mechanics/molecular mechanics/polarizable continuum model (QM/MM/PCM).

Methods

The atomic coordinates were obtained from the X-ray structures: PbRC from Rhodobacter sphaeroides at a 2.01-Å resolution (PDB code: 3I4D)32 and the PSII monomer unit (designated monomer A) of the PSII complexes from Thermosynechococcus vulcanus at a 1.9-Å resolution (PDB code: 3ARC).33 The energetics and electronic couplings were analyzed using the polarizable QM/MM/PCM method with the QuanPol code34 implemented in the GAMESS code,35 as in previous studies.36−39 The B3LYP functional and 6-31G* basis set were used for the QM region, whereas the polarizable amber-02 force field40 was used for the MM region. The PCM with a dielectric constant of 80 was used to reproduce the polarization of water molecules in the bulk region. The QM region was composed of quinones (QA and QB), the nonheme Fe complex (Fe, His-L190, His-L230, His-M219, His-M266, and Glu-M234), and the H-bond network of the distal carbonyl QB (Ser-L223, Asp-L213, Asp-M17, a water molecule that connects Asp-L213 and Asp-M17 (HOH-267, PDB code: 3I4D), and a water molecule that forms an H-bond network with Glu-M234 (HOH-308, PDB code: 3I4D(32))). The QM atoms were fully quantum-chemically optimized, whereas the MM atoms were fixed at certain positions in the crystal structure (PDB code: 3I4D).32 The protonation states of the amino acid residues were identical to those in previous studies;9 for example, Glu-L212 was protonated in the forward electron transfer from QA to QB.7,41−46

The overall electronic coupling between QA and QB (HQA–QB) was expressed using the framework of perturbation theory as follows:47,48

graphic file with name jp2c05713_m001.jpg 1.1
graphic file with name jp2c05713_m002.jpg 1.2
graphic file with name jp2c05713_m003.jpg 1.3

where HI is the superexchange coupling via the Ith intermediate state; VQA-I and VI-QB are the electronic couplings between the Ith intermediate state and QA and QB, respectively; ΔEI is the energy gap between the initial and intermediate states; E0 and EI are the energies of the initial and Ith intermediate states, respectively. The direct electronic coupling between QA and QB (VQA–QB) is negligible (QA···QB distance = ∼13 Å32). It should be noted that “superexchange coupling” presented here, i.e., for electron transfer via the virtual state,49 is not “superexchange coupling” for the strong antiferromagnetic coupling through a nonmagnetic anion (J-coupling,50−52 e.g., nonheme Fe complex in PbRC53 and PSII54 and Fe4S4 clusters in PSI55).

The electronic couplings, VQA–I and VI–QB, were evaluated by considering the Kohn–Sham matrix of the entire QM region and the molecular orbitals (MOs) of monomers56 in the QM/MM/PCM framework. Although the quantitative values depend on the functional, the B3LYP functional was used to evaluate the electronic coupling. [Fe(His)4Glu]+ is regarded as a single site for calculating the intermediate MOs, where the spin multiplicity is a quintet for the high-spin state of Fe2+ (S = 2). Apart from the chemical cores, all up- and down-spin MOs of [Fe(His)4Glu]+ are considered to be intermediate MOs. Electronic coupling was evaluated at the minimum energy conformation.

The energy difference between the initial and final states (ΔE) was evaluated based on QM/MM/PCM calculations of QA, QA•–, QB, and QB•– (eq 2), where the geometries of the monomers were optimized considering the charge-neutral and anion states.

graphic file with name jp2c05713_m004.jpg 2

The energy gap to the lowest reduced intermediate state was evaluated using QM/MM/PCM calculations of the monomers of QA and nonheme Fe complex as follows:

graphic file with name jp2c05713_m005.jpg 3.1

where Eunocci is the total energy with the ith unoccupied MO (lowest unoccpied MO, LUMO, when I= 1), the transfers of up- and down-spin electrons create [Fe(His)4Glu]0 with the multiplicity of sextet (S = 5/2) and quartet (S = 3/2), respectively. The energy gap via the ith unoccupied MO of the nonheme Fe complex was evaluated by considering the orbital energies, εunocci, as follows:

graphic file with name jp2c05713_m006.jpg 3.2

Using the same analogy, the energy gap to the lowest oxidized intermediate state was evaluated as follows:

graphic file with name jp2c05713_m007.jpg 4.1

where Eoccui is the total energy with the ith occupied MO (highest occupied MO, HOMO, when i = 1), the transfers of up- and down-spin electrons create [Fe(His)4Glu]2+ with the multiplicity of quartet (S = 3/2) and sextet (S = 5/2), respectively. The energy gap via the ith occupied MO of the nonheme Fe complex was evaluated as follows:

graphic file with name jp2c05713_m008.jpg 4.2

The electron transfer rate, kMLJ, was evaluated using the Marcus–Levich–Jortner theory57 (eq 5), which accounts for the Franck–Condon factor of discrete vibronic states of intramolecular modes and environmental reorganization energy.

graphic file with name jp2c05713_m009.jpg 5.1
graphic file with name jp2c05713_m010.jpg 5.2
graphic file with name jp2c05713_m011.jpg 5.3

where kMLJ consists of the electronic part, HQA–QB2, and intramolecular (FCintra) and environmental (FCenv) nuclear parts; ℏ is the Planck constant; FCintra (eq 5.2) is the overlap of vibrational wave functions between the initial and final states, which can be expressed as a product of the one-dimensional Franck–Condon factors along all intramolecular modes of QA and QB; and Si is the Huang–Rhys factor along the ith mode, which is expressed by the reorganization energy, λi, and frequency, ωi, as λi = Si × ℏωi. λi and ωi of QA and QB were calculated through geometry optimizations and normal-mode analysis using the QM/MM/PCM model, where the atomic displacements from the initial to final states were projected onto the normal modes. Subsequently, FCintra (eq 5.2) was evaluated considering the Huang–Rhys factor, Si.

FCenv (eq 5.3) is the thermally averaged Franck–Condon factor for the environment. kB and T are the Boltzmann constant and temperature, respectively. λenv is the reorganization energy of the environment, that is, the relaxation of all atoms in the crystal structure except for QA, QB, and the nonheme Fe complex, and ΔE is the bottom-to-bottom energy difference between the initial and final states. The sum of FCintra × FCenv for all possible vibrational excitations mi (i = 1 – n) was considered under the conditions of 0 ≥ ΔE + ∑miℏωi.

To evaluate λenv, the geometries of proteins, cofactors, and water molecules were optimized through classical MM calculations considering the atomic charges of QA and QB in the initial (QA•–···QB) and final (QA···QB•–) states, where the charge was determined using the restrained electrostatic potential (RESP) procedure.58

Results and Discussion

Electron Transfer in PbRC

The electronic coupling can be calculated for the LUMO between QA and QB via the MO of the nonheme Fe complex (i.e., superexchange coupling) because the direct electronic coupling between QA and QB (VQA–QB) is negligibly small due to the distance of >13 Å (Figure 1). The net superexchange coupling between QA and QB (HQA–QB) is 3.5–3.8 meV in PbRC (Table 1) when electron transfer occurs via the unoccupied MOs of the nonheme Fe complex (Figure 2a).

Figure 1.

Figure 1

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Electron transfer pathways for type-II reaction centers. (a) PbRC. (b) PSII. Red curved arrows indicate electron transfer. Black dotted lines indicate H bonds.

Table 1. Calculated Superexchange Coupling HQA–QB for Electron Transfer from QA to QB (meV).

  spin PbRC     PSII  
    native   ΔH2Oa nativeb formatec
(Ser)–OH partnerd   ···QB ···Asp–L213 ···QB ···QB ···QB
QA→Fe→QBe up –3.5 –1.1 –0.85 0.006 0.26
(unoccupied MO of Fef) down –3.8 –0.28 –0.42 0.009 –0.14
Fe→QB/QA→Feg up 0.17 –0.10 –0.12 0.21 0.20
(occupied MO of Fef) down 0.17 –0.021 0.54 0.18 0.39
  average –3.5 –0.72 –0.42 0.20 0.36
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a

In the absence of the water molecule adjacent to Glu-M234 (i.e., HOH-308, PDB code: 3I4D(32)).

b

See ref (39).

c

The hydroxyl group of bicarbonate was replaced with a methyl group (i.e., formate).

d

H-bond acceptor of serine (Ser-L223 in PbRC and D1-Ser264 in PSII).

e

Electron transfer via the virtual [Fe(His)4Glu]0 and [Fe(His)4HCO3–]0 intermediate states (Figure 2a).

f

Electron transfer via MOs of the nonheme Fe complex.

g

Electron transfer via the virtual [Fe(His)4Glu]2+ and [Fe(His)4HCO3–]2+ intermediate states (Figure 2b).

Figure 2.

Figure 2

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Superexchange pathways. (a) Electron transfer from QA to the nonheme Fe complex, followed by electron transfer from the nonheme Fe complex to QB (QA → Fe → QB), involving the reduced nonheme Fe complex ([Fe(His)4Glu]0) as an intermediate state. (b) Electron transfer from the nonheme Fe complex to QB, followed by electron transfer from QA to the nonheme Fe complex (Fe → QB/QA → Fe), involving the oxidized nonheme Fe complex ([Fe(His)4Glu]2+) as an intermediate state. [Fe(His)4Glu]0 and [Fe(His)4Glu]2+ are virtual states for superexchange and not thermally accessible.39

Remarkably, |HQA–QB| = 3.5 meV in PbRC (Table 1) is significantly larger than |HQA–QB| = 0.2 meV in PSII,39 despite the similar QA···QB distance of ∼13 Å for PbRC32 and PSII.33 The HQA–QB value for PbRC produces a time constant (1/kET) of 2–7 μs, assuming that ΔGET = −0.1 eV (e.g., 0.03–0.07 eV6,8,12−14) and λ = 0.8–0.9 eV (e.g., ref (59)). Using the same ΔGET and λ, the HQA–QB value for PSII produces the time constant of 140–390 μs. This result demonstrates that electron transfer from QA to QB is faster in PbRC than in PSII (Figure S2). The results are consistent with the experimentally measured 1/kET values in PbRC and PSII, where 1/kET ≪ 1/kAB(1) = 110 μs in PbRC11 and 1/kET = 200–400 μs in PSII,23 which suggests that HQA–QB is predominantly responsible for the difference in 1/kET. Below, we investigate the factors differentiating HQA–QB between the two type-II reaction centers.

(i) Cofactor arrangement. Electronic coupling is associated with the electron donor–acceptor distance. However, both the QA···QB distance (13 Å in the crystal structures for PbRC32 and PSII33) and the H-bond distance (e.g., His···QB distance = 2.7 Å in the QM/MM-optimized structures for PbRC9 and PSII18) are essentially the same in the two reaction centers.

(ii) Ligand carboxylic acid. PbRC has Glu-M234 as a ligand for Fe, whereas PSII has bicarbonate. The bicarbonate ligand plays a role in tuning the electron transfer from QA to QB in PSII.28,60 The electron paramagnetic resonance (EPR) experiments suggested that electron transfer was perturbed upon the replacement of bicarbonate with formate.61 However, the HQA–QB calculated by replacing bicarbonate with formate is similar to that in the wild-type PSII (Table 1). This is in line with the EPR experiment, that showed that the main effects of formate appeared on the QBH2 exchange29 (see below for further discussion).

(iii) Protein environment specific to each reaction center.

Remarkably, a water molecule, which donates an H-bond to the carboxyl O site of the Glu-M234 ligand (Glu-M234···water = 2.6 Å32), contributes significantly to HQA–QB in PbRC. This is indicated by the MO of the nonheme Fe complex that is delocalized over the water molecule and thus migrated toward QB (Figure 3). |HQA–QB| significantly decreases to 0.42 meV once the water molecule is depleted (Table 1, Figure S3), which indicates that the water molecule adjacent to Glu-M234 is responsible for the especially large HQA–QB in PbRC (see below for other factors).

Figure 3.

Figure 3

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Unoccupied MOs of the intermediate nonheme Fe complex contributing to a strong coupling via H2O. For clarity, the protein electrostatic environment is not shown.

The water molecule exists in the QB binding pocket, regardless of the presence (e.g., HOH-M308, PDB code: 3I4D)32 or absence (e.g., HOH-M1067, PDB code: 1L9B;62 HOH-M584, PDB code: 7P2C(63)) of QB in the PbRC crystal structures. In particular, the water molecule is well ordered, as indicated by the lowest B-factor value (= 32) among all water molecules in the QB binding pocket.32 It seems plausible that the binding of the water molecule is stabilized by the negatively charged carboxyl O atom of Glu-M234. In response to the loss of QBH2, the water molecule is involved in the H-bond network with Glu-L212 and His-L190,62 leading to the reprotonation of deprotonated His-L190 via protonated Glu-L212.9 The corresponding water molecule is absent at the QA site, because neither water molecule nor water channel exists in the QA binding pocket.

In contrast, the existence of a corresponding water molecule at the bicarbonate moiety is inhibited by the bulky side-chain of D1-Tyr246 (bicarbonate···D1-Tyr246 = 3.2 Å33), which is conserved even in the earliest evolving D1 proteins.64 The role of the water molecule in superexchange electron tunneling cannot be substituted by D1-Tyr246, because D1-Tyr246 does not form an H-bond with the bicarbonate O site (unless the bidentate to monodentate reorientation of the bicarbonate ligand occurs65). As D1-Tyr246 is located in the de-loop region, which is absent in PbRC but crucial to PSII functioning (e.g., photoinhibition or light susceptibility66−68), the low HQA–QB value due to the loss of water molecules is a distinct feature of PSII.

The D1-Y246A mutation may provide room for the binding of a water molecule to the bicarbonate moiety. However, the D1-Y246A mutant exhibited sensitivity to high light, rapidly losing its oxygen evolving capacity in high light.68 In addition, the D1-Y246A mutant lost its ability to grow photoautotrophically.68 Furthermore, electron transfer from QA to QB was impaired in the D1-Y246A cells, which suggests that structural changes occurred upon the mutation.68 Thus, D1-Tyr246 is a prerequisite for PSII functioning, even if the water binding loss decreases HQA–QB.

Conformational Gating

The rate of the first electron transfer from QA to QB is independent of the driving force due to conformational gating.11 Based on the observations that (i) the light-exposed crystal structure shows that QB forms an H-bond with His-L190;69 (ii) QB remains at the unique binding position during electron transfer;70−72 and (iii) Ser-L223, which forms an H-bond with Asp-L213 in charge neutral QB, reorients toward the QB carbonyl O site in QB•–;7,8,73,74 the formation of an H-bond between Ser-L223 and QB is likely to play a significant role in conformational gating.

HQA–QB increases significantly from −0.72 to −3.5 meV as the Ser-L223···QB H-bond forms (Table 1). The observed increase in HQA–QB is due to the elongation of the H-bond network at the QB site. The H-bond network QA···His-M219···Fe···His-L190···QB is extended to QA···His-M219···Fe···His-L190···QB···Ser-L223···Asp-L213···H2O···Asp-M17 in response to the formation of the Ser-L223···QB H-bond,9 thus facilitating the charge delocalization over the QB site (Figure 4).

Figure 4.

Figure 4

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Insight into the conformational gating. (Top) Representative MOs of QA and QB hybridized with σ* orbitals of the histidine ligands in the nonheme Fe complex. (Middle) Spectra of superexchange coupling (HI) for the QA-to-QB electron tunneling via the unoccupied MOs of the nonheme Fe complex in PbRC: up-spin. The horizontal axis indicates the energy gap (ΔEI). See Figure S4 for down-spin. (Bottom) Overview. The formation of the Ser-L223···QB H-bond increases HQA–QB significantly, facilitating the superexchange tunneling between QA and QB. Yellow backgrounds indicate the magnitude of the superexchange coupling along the delocalized H-bond network. The red circle indicates the contribution of the water molecule at the Glu-M234 moiety to HQA–QB.

QAside. The formation of the H-bond network stabilizes the MOs of QB, altering the interacting MOs of the nonheme Fe complex. The delocalization of MOs over the His-M219 moiety at the QA site is more pronounced in response to the formation of the H-bond network at the QB site (Figure 4, top), contributing to an increase in |HQA-QB|.

QBside. The hybridized σ and σ* orbitals of the histidine ligands are strongly coupled with the LUMOs of QA and QB (π orbits) and enhance the electronic coupling in PbRC (Figure 4, top). Thus, delocalized histidine σ and σ* orbitals provided an efficient route for superexchange electron tunneling.39,75 These features are commonly observed in both PSII39 and PbRC. Remarkably, the His-L190···QB interaction shows that the contributions of the histidine π and π* orbitals are pronounced before the formation of the H-bond network, whereas those of σ and σ* orbitals are pronounced after the formation of the H-bond network (Figure 4, top). The contributions of the histidine σ and σ* orbitals to the superexchange coupling are larger than those of π and π* orbitals.39 Thus, the conformational gating is likely to enhance the superexchange electron tunneling by switching the histidine π and π* to σ and σ* orbitals in the His-L190···QB interaction (Figure 4, top).

These indicate that the H-bond formation is responsible not only for stabilizing the QB•– state and increasing the driving force8,73 but also for enhancing the superexchange coupling, that is, electron transfer gating (Figure 4).

Discrepancy in the Electron Transfer Mechanism between PbRC and PSII

HQA–QB is significantly larger for electron transfer via unoccupied MOs of the nonheme Fe complex than via occupied MOs of the nonheme Fe complex in PbRC (Table 1), whereas HQA–QB is significantly smaller for electron transfer via unoccupied MOs of the nonheme Fe complex than via occupied MOs of the nonheme Fe complex in PSII.39 That is, in the superexchange electron tunneling, the following characteristic is pronounced: the QA-to-Fe electron transfer is followed by the Fe-to-QB electron transfer (QA → Fe → QB) in PbRC (Figure 2a), whereas the Fe-to-QB electron transfer is followed by the QA-to-Fe electron transfer (Fe → QB/QA → Fe) in PSII (Figure 2b). Note that [Fe(His)4HCO3–]0 and [Fe(His)4HCO3–]2+ are virtual intermediate states for superexchange and not thermally accessible.39 Consistently, the release of an electron from the nonheme Fe complex occurs in PSII,76 as indicated by the experimentally measured value of Em(Fe2+/Fe3+) = 400 mV.30,31 In contrast, the release of an electron from the nonheme Fe complex is energetically unfavorable in PbRC. Time-resolved X-ray absorption experiments also suggested that the Fe2+ to Fe3+ oxidation was not observed during the electron transfer from QA to QB in PbRC.77 As suggested previously,77,78 the present result does not support the Fe → QB/QA → Fe electron transfer in PbRC proposed by Remy and Gerwert.70

All of the cases presented here show that the electron transfer route via the occupied MOs of the nonheme Fe complex (Fe → QB/QA → Fe) is commonly ∼10 times less advantageous than the route via the unoccupied MOs (QA → Fe → QB) in terms of superexchange coupling (Table 1). Therefore, the utilization of the electron transfer route via the unoccupied MOs of the nonheme Fe complex is considered a strategy to enhance superexchange coupling and facilitate electron transfer between QA and QB in type-II reaction centers. The superexchange coupling for electron transfer via occupied MOs of the nonheme Fe complex is similar in PbRC and PSII (0.2–0.3 meV, Table 1); the discrepancy is only pronounced in the superexchange coupling for electron transfer via the unoccupied MOs of the nonheme Fe complex (3–4 meV in PbRC and 0.01 meV in PSII, Table 1).

Intriguingly, superexchange coupling via the unoccupied MOs of the nonheme Fe complex is 10 times larger in formate-PSII than in the native bicarbonate-PSII (Table 1). As HQA–QB is larger in formate-PSII than in native bicarbonate-PSII, the inhibition of the electron transfer process (i.e., kET) is unlikely the main effect of bicarbonate being replaced with formate, which is in line with the EPR studies that suggested that formate mainly affected the QBH2 exchange.29 These results suggest that the bicarbonate ligand specifically inhibits the efficient electron transfer route via the unoccupied MOs of the nonheme Fe complex because the virtual [Fe(His)4HCO3–]0 state is destabilized by −Oδ−H near the −COO– group. This characteristic can also be understood by Em(Fe2+/3+: formate) ≫ Em(Fe2+/3+: bicarbonate) in PSII,79 which indicates that the bicarbonate ligand preferentially facilitates the oxidation of the nonheme Fe complex rather than its reduction. It seems likely that PSII is only allowed to use the less efficient electron transfer route via the occupied MOs of the nonheme Fe complex. This could explain why electron transfer from QA to QB is slower in PSII than in PbRC, despite the conserved cofactor arrangement.

Conclusions

The net superexchange coupling between QA and QB is significantly larger in PbRC than in PSII (Table 1), irrespective of the conserved cofactor arrangement (Figure 1). This is primarily due to the presence of the water molecule that donates an H bond to the Glu-M234 ligand of the nonheme Fe complex in PbRC and secondarily due to the presence of the bicarbonate ligand in PSII. In PbRC, the MO of the nonheme Fe complex is delocalized over the water molecule and thus migrates toward QB (Figures 4 and S2). The formation of the H bond between QB and Ser-L223 and the extension of the H-bond network at the QB site7,8,73,74 induce the transformation of the π to σ interaction between the histidine ligand and the quinone, enhancing the superexchange electron tunneling, HQA-QB, serving as “a conformational gating11” (Figure 4).

The corresponding water molecule cannot exist because of D1-Tyr246 near the bicarbonate ligand of the nonheme Fe complex in PSII. The electron transfer route via the unoccupied MOs of the nonheme Fe complex (QA → Fe → QB via the virtual [Fe(His)4Glu]0 intermediate state) is ∼10 times more efficient than the route via the occupied MOs of the nonheme Fe complex (Fe → QB/QA → Fe via the virtual [Fe(His)4Glu]2+ intermediate state; Table 1). PbRC can use this efficient route via the unoccupied MOs of the nonheme Fe complex. However, the bicarbonate ligand specifically inhibits the electron transfer route via the unoccupied MOs of the nonheme Fe complex in PSII; this may explain (i) why the electron transfer is faster in PbRC than in PSII and (ii) why the nonheme Fe complex is redox-active in PSII but not in PbRC. It seems possible that the utilization of the electron transfer route via the unoccupied MOs of the nonheme Fe complex is an advantageous strategy to enhance the superexchange coupling and thus facilitate electron transfer between QA and QB in type-II reaction centers (e.g., PbRC). This strategy may simultaneously lose the redox activity of the nonheme Fe complex (e.g., PbRC). These are the bases of the role of the nonheme Fe complex in type-II reaction centers, gating the electron transfer and finalizing the photoinduced charge separation.

Acknowledgments

This research was supported by JSPS KAKENHI (JP20H05090 to H.I.) and Interdisciplinary Computational Science Program in CCS, University of Tsukuba.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.jpcb.2c05713.

  • Molecular structures of quinones; time constant of the QA → QB electron tunneling based on the Marcus–Levich–Jortner theory; spectra of superexchange coupling for the QA-to-QB electron tunneling with respect to the energy gap in the H2O-depleted PbRC; spectra of superexchange coupling for the QA-to-QB electron tunneling via the unoccupied MOs of the nonheme Fe complex for down-spin (PDF)

Author Contributions

§ These authors contributed equally to this work.

The authors declare no competing financial interest.

Supplementary Material

jp2c05713_si_001.pdf (294KB, pdf)

References

  1. Michel H.; Deisenhofer J. Relevance of the photosynthetic reaction center from purple bacteria to the structure of photosystem II. Biochemistry 1988, 27, 1–7. 10.1021/bi00401a001. [DOI] [Google Scholar]
  2. Rutherford A. W.; Faller P. Photosystem II: evolutionary perspectives. Philos. Trans. R. Soc. London B 2003, 358, 245–253. 10.1098/rstb.2002.1186. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Cardona T.; Sedoud A.; Cox N.; Rutherford A. W. Charge separation in photosystem II: a comparative and evolutionary overview. Biochim. Biophys. Acta 2012, 1817, 26–43. 10.1016/j.bbabio.2011.07.012. [DOI] [PubMed] [Google Scholar]
  4. Parson W. W.; Chu Z.-T.; Warshel A. Electrostatic control of charge separation in bacterial photosynthesis. Biochim. Biophys. Acta 1990, 1017, 251–272. 10.1016/0005-2728(90)90192-7. [DOI] [PubMed] [Google Scholar]
  5. Steffen M. A.; Lao K.; Boxer S. G. Dielectric asymmetry in the photosynthetic reaction center. Science 1994, 264, 810–6. 10.1126/science.264.5160.810. [DOI] [PubMed] [Google Scholar]
  6. Okamura M. Y.; Paddock M. L.; Graige M. S.; Feher G. Proton and electron transfer in bacterial reaction centers. Biochim. Biophys. Acta 2000, 1458, 148–163. 10.1016/S0005-2728(00)00065-7. [DOI] [PubMed] [Google Scholar]
  7. Alexov E. G.; Gunner M. R. Calculated protein and proton motions coupled to electron transfer: electron transfer from QA– to QB in bacterial photosynthetic reaction centers. Biochemistry 1999, 38, 8253–8270. 10.1021/bi982700a. [DOI] [PubMed] [Google Scholar]
  8. Ishikita H.; Knapp E.-W. Variation of Ser-L223 hydrogen bonding with the QB redox state in reaction centers from Rhodobacter sphaeroides. J. Am. Chem. Soc. 2004, 126, 8059–8064. 10.1021/ja038092q. [DOI] [PubMed] [Google Scholar]
  9. Sugo Y.; Saito K.; Ishikita H. Mechanism of the formation of proton transfer pathways in photosynthetic reaction centers. Proc. Natl. Acad. Sci. U. S. A. 2021, 118, e2103203118. 10.1073/pnas.2103203118. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Osváth S.; Maróti P. Coupling of cytochrome and quinone turnovers in the photocycle of reaction centers from the photosynthetic bacterium Rhodobacter sphaeroides. Biophys. J. 1997, 73, 972–982. 10.1016/S0006-3495(97)78130-X. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Graige M. S.; Feher G.; Okamura M. Y. Conformational gating of the electron transfer reaction QA–QB → QAQB– in bacterial reaction centers of Rhodobacter sphaeroides determined by a driving force assay. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 11679–11684. 10.1073/pnas.95.20.11679. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Li J.; Takahashi E.; Gunner M. R. –ΔGoAB and pH dependence of the electron transfer from P+QA–QB to P+QAQB– in Rhodobacter sphaeroides reaction centers. Biochemistry 2000, 39, 7445–7454. 10.1021/bi992591f. [DOI] [PubMed] [Google Scholar]
  13. Alexov E.; Miksovska J.; Baciou L.; Schiffer M.; Hanson D. K.; Sebban P.; Gunner M. R. Modeling the effects of mutations on the free energy of the first electron transfer from QA– to QB in photosynthetic reaction centers. Biochemistry 2000, 39, 5940–5952. 10.1021/bi9929498. [DOI] [PubMed] [Google Scholar]
  14. Zhu Z.; Gunner M. R. Energetics of quinone-dependent electron and proton transfers in Rhodobacter sphaeroides photosynthetic reaction centers. Biochemistry 2005, 44, 82–96. 10.1021/bi048348k. [DOI] [PubMed] [Google Scholar]
  15. Muh F.; Glockner C.; Hellmich J.; Zouni A. Light-induced quinone reduction in photosystem II. Biochim. Biophys. Acta 2012, 1817, 44–65. 10.1016/j.bbabio.2011.05.021. [DOI] [PubMed] [Google Scholar]
  16. Debus R. J.; Feher G.; Okamura M. Y. Iron-depleted reaction centers from Rhodopseudomonas sphaeroides R-26.1: characterization and reconstitution with Fe2+, Mn2+, Co2+, Ni2+, Cu2+, and Zn2+. Biochemistry 1986, 25, 2276–2287. 10.1021/bi00356a064. [DOI] [PubMed] [Google Scholar]
  17. Ishikita H.; Knapp E.-W. Electrostatic role of the non-heme iron complex in bacterial photosynthetic reaction center. FEBS Lett. 2006, 580, 4567–4570. 10.1016/j.febslet.2006.07.023. [DOI] [PubMed] [Google Scholar]
  18. Saito K.; Rutherford A. W.; Ishikita H. Mechanism of proton-coupled quinone reduction in Photosystem II. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 954–959. 10.1073/pnas.1212957110. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Saito K.; Mandal M.; Ishikita H. Redox potentials along the redox-active low-barrier H-bonds in electron transfer pathways. Phys. Chem. Chem. Phys. 2020, 22, 25467–25473. 10.1039/D0CP04265J. [DOI] [PubMed] [Google Scholar]
  20. Hördt A.; López M. G.; Meier-Kolthoff J. P.; Schleuning M.; Weinhold L.-M.; Tindall B. J.; Gronow S.; Kyrpides N. C.; Woyke T.; Göker M., Analysis of 1,000+ type-strain genomes substantially improves taxonomic classification of Alphaproteobacteria. Front. Microbiol. 2020, 11, 10.3389/fmicb.2020.00468. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Kato Y.; Nagao R.; Noguchi T. Redox potential of the terminal quinone electron acceptor QB in photosystem II reveals the mechanism of electron transfer regulation. Proc. Natl. Acad. Sci. U. S. A. 2016, 113, 620–5. 10.1073/pnas.1520211113. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. De Causmaecker S.; Douglass J. S.; Fantuzzi A.; Nitschke W.; Rutherford A. W. Energetics of the exchangeable quinone, QB, in Photosystem II. Proc. Natl. Acad. Sci. U. S. A. 2019, 116, 19458–19463. 10.1073/pnas.1910675116. [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. de Wijn R.; van Gorkom H. J. Kinetics of electron transfer from QA to QB in photosystem II. Biochemistry 2001, 40, 11912–11922. 10.1021/bi010852r. [DOI] [PubMed] [Google Scholar]
  24. Velthuys B. R. Electron-dependent competition between plastoquinone and inhibitors for binding to photosystem II. FEBS Lett. 1981, 126, 277–281. 10.1016/0014-5793(81)80260-8. [DOI] [Google Scholar]
  25. Naber J. D.; van Rensen J. J. S.. Determination of the exchange parameters of herbicides on the QB-protein of photosystem II. In Progress in Photosynthesis Research: Vol. 3 Proceedings of the VIIth International Congress on Photosynthesis; Providence, Rhode Island, August 10–15, 1986; Biggins J., Ed.; Springer: Dordrecht, The Netherlands, 1987; pp 767–770. [Google Scholar]
  26. Sugo Y.; Saito K.; Ishikita H. Conformational changes and H-bond rearrangements during quinone release in photosystem II. Biochemistry 2022, 61, 1836–1843. 10.1021/acs.biochem.2c00324. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Jursinic P.; Warden J.; Govindjee A major site of bicarbonate effect in system II reaction. Evidence from ESR signal IIvf, fast fluorescence yield changes and delayed light emission. Biochim. Biophys. Acta 1976, 440, 322–330. 10.1016/0005-2728(76)90066-9. [DOI] [PubMed] [Google Scholar]
  28. Eaton-Rye J. J.; Govindjee Electron transfer through the quinone acceptor complex of photosystem II after one or two actinic flashes in bicarbonate-depleted spinach thylakoid membranes. Biochim. Biophys. Acta 1988, 935, 248–257. 10.1016/0005-2728(88)90221-6. [DOI] [Google Scholar]
  29. Sedoud A.; Kastner L.; Cox N.; El-Alaoui S.; Kirilovsky D.; Rutherford A. W. Effects of formate binding on the quinone-iron electron acceptor complex of photosystem II. Biochim. Biophys. Acta 2011, 1807, 216–26. 10.1016/j.bbabio.2010.10.019. [DOI] [PubMed] [Google Scholar]
  30. Bowes J. M.; Crofts A. R.; Itoh S. A high potential acceptor for Photosystem II. Biochim. Biophys. Acta 1979, 547, 320–335. 10.1016/0005-2728(79)90014-8. [DOI] [PubMed] [Google Scholar]
  31. Wraight C. A. Modulation of herbicide-binding by the redox state of Q400, an endogenous component of photosystem II. Biochim. Biophys. Acta 1985, 809, 320–330. 10.1016/0005-2728(85)90181-1. [DOI] [Google Scholar]
  32. Roszak A. W.; Moulisova V.; Reksodipuro A. D. P.; Gardiner A. T.; Fujii R.; Hashimoto H.; Isaacs N. W.; Cogdell R. J. New insights into the structure of the reaction centre from Blastochloris viridis: evolution in the laboratory. Biochem. J. 2012, 442, 27–37. 10.1042/BJ20111540. [DOI] [PubMed] [Google Scholar]
  33. Umena Y.; Kawakami K.; Shen J.-R.; Kamiya N. Crystal structure of oxygen-evolving photosystem II at a resolution of 1.9 Å. Nature 2011, 473, 55–60. 10.1038/nature09913. [DOI] [PubMed] [Google Scholar]
  34. Thellamurege N. M.; Si D. J.; Cui F. C.; Zhu H. B.; Lai R.; Li H. QuanPol: a full spectrum and seamless QM/MM program. J. Comput. Chem. 2013, 34, 2816–2833. 10.1002/jcc.23435. [DOI] [PubMed] [Google Scholar]
  35. Schmidt M. W.; et al. General atomic and molecular electronic-structure system. J. Comput. Chem. 1993, 14, 1347–1363. 10.1002/jcc.540141112. [DOI] [Google Scholar]
  36. Tamura H.; Saito K.; Ishikita H. Acquirement of water-splitting ability and alteration of the charge-separation mechanism in photosynthetic reaction centers. Proc. Natl. Acad. Sci. U. S. A. 2020, 117, 16373–16382. 10.1073/pnas.2000895117. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Mitsuhashi K.; Tamura H.; Saito K.; Ishikita H. Nature of asymmetric electron transfer in the symmetric pathways of photosystem I. J. Phys. Chem. B 2021, 125, 2879–2885. 10.1021/acs.jpcb.0c10885. [DOI] [PubMed] [Google Scholar]
  38. Tamura H.; Saito K.; Ishikita H. The origin of unidirectional charge separation in photosynthetic reaction centers: nonadiabatic quantum dynamics of exciton and charge in pigment–protein complexes. Chem. Sci. 2021, 12, 8131–8140. 10.1039/D1SC01497H. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Tamura H.; Saito K.; Ishikita H. Long-range electron tunneling from the primary to secondary quinones in photosystem II enhanced by hydrogen bonds with a nonheme Fe complex. J. Phys. Chem. B 2021, 125, 13460–13466. 10.1021/acs.jpcb.1c09538. [DOI] [PubMed] [Google Scholar]
  40. Cieplak P.; Caldwell J.; Kollman P. Molecular mechanical models for organic and biological systems going beyond the atom centered two body additive approximation: Aqueous solution free energies of methanol and N-methyl acetamide, nucleic acid base, and amide hydrogen bonding and chloroform/water partition coefficients of the nucleic acid bases. J. Comput. Chem. 2001, 22, 1048–1057. 10.1002/jcc.1065. [DOI] [Google Scholar]
  41. Maróti P.; Wraight C. W. Flash-induced H+ binding by bacterial photosynthetic reaction centers: influences of the redox states of the acceptor quinones and primary donor. Biochim. Biophys. Acta 1988, 934, 329–347. 10.1016/0005-2728(88)90092-8. [DOI] [Google Scholar]
  42. McPherson P. H.; Nagarajan V.; Parson W. W.; Okamura M. Y.; Feher G. pH-dependence of the free energy gap between DQA and D+QA– determined from delayed fluorescence in reaction centers from Rhodobacter sphaeroides R-26. Biochim. Biophys. Acta 1990, 1019, 91–94. 10.1016/0005-2728(90)90128-Q. [DOI] [Google Scholar]
  43. Hienerwadel R.; Grzybek S.; Fogel C.; Kreutz W.; Okamura M. Y.; Paddock M. L.; Breton J.; Nabedryk E.; Maentele W. Protonation of Glu L212 following QB– formation in the photosynthetic reaction center of Rhodobacter sphaeroides: evidence from time-resolved infrared spectroscopy. Biochemistry 1995, 34, 2832–2843. 10.1021/bi00009a013. [DOI] [PubMed] [Google Scholar]
  44. Nabedryk E.; Breton J.; Hienerwadel R.; Fogel C.; Maentele W.; Paddock M. L.; Okamura M. Y. Fourier transform infrared difference spectroscopy of secondary quinone acceptor photoreduction in proton transfer mutants of Rhodobacter sphaeroides. Biochemistry 1995, 34, 14722–14732. 10.1021/bi00045a013. [DOI] [PubMed] [Google Scholar]
  45. Miksovska J.; Maróti P.; Tandori J.; Schiffer M.; Hanson D. K.; Sebban P. Distant electrostatic interactions modulate the free energy level of QA– in the photosynthetic reaction center. Biochemistry 1996, 35, 15411–15417. 10.1021/bi961299u. [DOI] [PubMed] [Google Scholar]
  46. Ishikita H.; Morra G.; Knapp E.-W. Redox potential of quinones in photosynthetic reaction centers from Rhodobacter sphaeroides: dependence on protonation of Glu-L212 and Asp-L213. Biochemistry 2003, 42, 3882–3892. 10.1021/bi026781t. [DOI] [PubMed] [Google Scholar]
  47. Todd M. D.; Nitzan A.; Ratner M. A. Electron transfer via superexchange: a time-dependent approach. J. Phys. Chem. 1993, 97, 29–33. 10.1021/j100103a008. [DOI] [Google Scholar]
  48. Ito H.; Nakatsuji H. Roles of proteins in the electron transfer in the photosynthetic reaction center of Rhodopseudomonas viridis: bacteriopheophytin to ubiquinone. J. Comput. Chem. 2001, 22, 265–272. . [DOI] [Google Scholar]
  49. Natali M.; Campagna S.; Scandola F. Photoinduced electron transfer across molecular bridges: electron- and hole-transfer superexchange pathways. Chem. Soc. Rev. 2014, 43, 4005–4018. 10.1039/C3CS60463B. [DOI] [PubMed] [Google Scholar]
  50. Anderson P. W. Antiferromagnetism. Theory of superexchange interaction. Phys. Rev. 1950, 79, 350–356. 10.1103/PhysRev.79.350. [DOI] [Google Scholar]
  51. Goodenough J. B. Theory of the role of covalence in the perovskite-type manganites [La, M(II)]MnO3. Phys. Rev. 1955, 100, 564–573. 10.1103/PhysRev.100.564. [DOI] [Google Scholar]
  52. Kanamori J. Superexchange interaction and symmetry properties of electron orbitals. J. Phys. Chem. Solids 1959, 10, 87–98. 10.1016/0022-3697(59)90061-7. [DOI] [Google Scholar]
  53. Butler W. F.; Calvo R.; Fredkin D. R.; Isaacson R. A.; Okamura M. Y.; Feher G. The electronic structure of Fe2+ in reaction centers from Rhodopseudomonas sphaeroides. III. EPR measurements of the reduced acceptor complex. Biophys. J. 1984, 45, 947–73. 10.1016/S0006-3495(84)84241-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
  54. Cox N.; Jin L.; Jaszewski A.; Smith P. J.; Krausz E.; Rutherford A. W.; Pace R. The semiquinone-iron complex of photosystem II: structural insights from ESR and theoretical simulation; evidence that the native ligand to the non-heme iron is carbonate. Biophys. J. 2009, 97, 2024–33. 10.1016/j.bpj.2009.06.033. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Kanda T.; Saito K.; Ishikita H. Mechanism of mixed-valence Fe2.5+···Fe2.5+ formation in Fe4S4 clusters in the ferredoxin binding motif. J. Phys. Chem. B 2022, 126, 3059–3066. 10.1021/acs.jpcb.2c01320. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Tamura H.; et al. Theoretical analysis on the optoelectronic properties of single crystals of thiophene-furan-phenylene co-oligomers: efficient photoluminescence due to molecular bending. J. Phys. Chem. C 2013, 117, 8072–8078. 10.1021/jp400646n. [DOI] [Google Scholar]
  57. Marcus R. A. Electron transfer reactions in chemistry. Theory and experiment. Rev. Mod. Phys. 1993, 65, 599–610. 10.1103/RevModPhys.65.599. [DOI] [Google Scholar]
  58. Bayly C. I.; Cieplak P.; Cornell W. D.; Kollman P. A. A well-behaved electrostatic potential based method using charge restraints for deriving atomic charges: the RESP model. J. Phys. Chem. 1993, 97, 10269–10280. 10.1021/j100142a004. [DOI] [Google Scholar]
  59. Page C. C.; Moser C. C.; Chen X.; Dutton P. L. Natural engineering principles of electron tunnelling in biological oxidation-reduction. Nature 1999, 402, 47–52. 10.1038/46972. [DOI] [PubMed] [Google Scholar]
  60. Eaton-Rye J. J.; Govindjee Electron transfer through the quinone acceptor complex of photosystem II in bicarbonate-depleted spinach thylakoid membranes as a function of actinic flash number and frequency. Biochim. Biophys. Acta 1988, 935, 237–247. 10.1016/0005-2728(88)90220-4. [DOI] [Google Scholar]
  61. Vermaas W. F. J.; Rutherford A. W. EPR measurements on the effects of bicarbonate and triazine resistance on the acceptor side of photosystem II. FEBS Lett. 1984, 175, 243–248. 10.1016/0014-5793(84)80744-9. [DOI] [Google Scholar]
  62. Axelrod H. L.; Abresch E. C.; Okamura M. Y.; Yeh A. P.; Rees D. C.; Feher G. X-ray structure determination of the cytochrome c2: reaction center electron transfer complex from Rhodobacter sphaeroides. J. Mol. Biol. 2002, 319, 501–515. 10.1016/S0022-2836(02)00168-7. [DOI] [PubMed] [Google Scholar]
  63. Selikhanov G.; Fufina T.; Guenther S.; Meents A.; Gabdulkhakov A.; Vasilieva L. X-ray structure of the Rhodobacter sphaeroides reaction center with an M197 Phe→His substitution clarifies the properties of the mutant complex. IUCrJ. 2022, 9, 261–271. 10.1107/S2052252521013178. [DOI] [PMC free article] [PubMed] [Google Scholar]
  64. Cardona T.; Murray J. W.; Rutherford A. W. Origin and evolution of water oxidation before the last common ancestor of the cyanobacteria. Mol. Biol. Evol. 2015, 32, 1310–1328. 10.1093/molbev/msv024. [DOI] [PMC free article] [PubMed] [Google Scholar]
  65. Sugo Y.; Ishikita H.. Proton-mediated photoprotection mechanism in photosystem II. Front. Plant Sci. 2022, 13, 10.3389/fpls.2022.934736. [DOI] [PMC free article] [PubMed] [Google Scholar]
  66. Maenpaa P.; Miranda T.; Tyystjarvi E.; Tyystjarvi T.; Govindjee; Ducruet J. M.; Etienne A. L.; Kirilovsky D. A mutation in the D-de loop of D1 modifies the stability of the S2QA– and S2QB– states in photosystem II. Plant Physiol. 1995, 107, 187–197. 10.1104/pp.107.1.187. [DOI] [PMC free article] [PubMed] [Google Scholar]
  67. Kless H.; Oren-Shamir M.; Malkin S.; McIntosh L.; Edelman M. The D-E region of the D1 protein is involved in multiple quinone and herbicide interactions in photosystem II. Biochemistry 1994, 33, 10501–7. 10.1021/bi00200a035. [DOI] [PubMed] [Google Scholar]
  68. Forsman J. A.; Vass I.; Eaton-Rye J. J. D1:Glu244 and D1:Tyr246 of the bicarbonate-binding environment of Photosystem II moderate high light susceptibility and electron transfer through the quinone-Fe-acceptor complex. Biochim. Biophys. Acta 2019, 1860, 148054. 10.1016/j.bbabio.2019.07.009. [DOI] [PubMed] [Google Scholar]
  69. Stowell M. H. B.; McPhillips T. M.; Rees D. C.; Soltis S. M.; Abresch E.; Feher G. Light-induced structural changes in photosynthetic reaction center: implications for mechanism of electron-proton transfer. Science 1997, 276, 812–816. 10.1126/science.276.5313.812. [DOI] [PubMed] [Google Scholar]
  70. Remy A.; Gerwert K. Coupling of light-induced electron transfer to proton uptake in photosynthesis. Nat. Struct. Biol. 2003, 10, 637–644. 10.1038/nsb954. [DOI] [PubMed] [Google Scholar]
  71. Breton J. Absence of large-scale displacement of quinone QB in bacterial photosynthetic reaction centers. Biochemistry 2004, 43, 3318–3326. 10.1021/bi049811w. [DOI] [PubMed] [Google Scholar]
  72. Baxter R. H. G.; Ponomarenko N.; Srajer V.; Pahl R.; Moffat K.; Norris J. R. Time-resolved crystallographic studies of light-induced structural changes in the photosynthetic reaction center. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 5982–5987. 10.1073/pnas.0306840101. [DOI] [PMC free article] [PubMed] [Google Scholar]
  73. Ishikita H.; Knapp E.-W. Control of quinone redox potentials in photosystem II: electron transfer and photoprotection. J. Am. Chem. Soc. 2005, 127, 14714–14720. 10.1021/ja052567r. [DOI] [PubMed] [Google Scholar]
  74. Paddock M. L.; Flores M.; Isaacson R.; Chang C.; Abresch E. C.; Okamura M. Y. ENDOR spectroscopy reveals light induced movement of the H-bond from Ser-L223 upon forming the semiquinone (QB–•) in reaction centers from Rhodobacter sphaeroides. Biochemistry 2007, 46, 8234–43. 10.1021/bi7005256. [DOI] [PMC free article] [PubMed] [Google Scholar]
  75. Balabin I. A.; Onuchic J. N. Dynamically controlled protein tunneling paths in photosynthetic reaction centers. Science 2000, 290, 114–7. 10.1126/science.290.5489.114. [DOI] [PubMed] [Google Scholar]
  76. Petrouleas V.; Diner B. A. Identification of Q400, a high-potential electron acceptor of Photosystem II, with the iron of the quinone-iron acceptor complex. Biochim. Biophys. Acta 1986, 849, 264–275. 10.1016/0005-2728(86)90033-2. [DOI] [Google Scholar]
  77. Hermes S.; Bremm O.; Garczarek F.; Derrien V.; Liebisch P.; Loja P.; Sebban P.; Gerwert K.; Haumann M. A time-resolved iron-specific X-ray absorption experiment yields no evidence for an Fe2+ → Fe3+ transition during QA– → QB electron transfer in the photosynthetic reaction center. Biochemistry 2006, 45, 353–359. 10.1021/bi0515725. [DOI] [PubMed] [Google Scholar]
  78. Nabedryk E.; Breton J. Coupling of electron transfer to proton uptake at the QB site of the bacterial reaction center: A perspective from FTIR difference spectroscopy. Biochim. Biophys. Acta 2008, 1777, 1229–1248. 10.1016/j.bbabio.2008.06.012. [DOI] [PubMed] [Google Scholar]
  79. Deligiannakis Y.; Petrouleas V.; Diner B. A. Binding of carboxylate anions at the non-heme Fe(II) of PSII. I. Effects on the QA–Fe2+ and QAFe3+ EPR spectra and the redox properties of the iron. Biochim. Biophys. Acta 1994, 1188, 260–270. 10.1016/0005-2728(94)90044-2. [DOI] [Google Scholar]

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