Proton coupled electron transfer and redox active tyrosines in Photosystem II

Abstract

In this article, progress in understanding proton coupled electron transfer (PCET) in photosystem II is reviewed. Changes in acidity/basicity may accompany oxidation/reduction reactions in biological catalysis. Alterations in the proton transfer pathway can then be used to alter the rates of the electron transfer reactions. Studies of the bioenergetic complexes have played a central role in advancing our understanding of PCET. Because oxidation of the tyrosine results in deprotonation of the phenolic oxygen, redox active tyrosines are involved in PCET reactions in several enzymes. This review focuses on PCET involving the redox active tyrosines in photosystem II. Photosystem II catalyzes the light-driven oxidation of water and reduction of plastoquinone. Photosystem II provides a paradigm for the study of redox active tyrosines, because this photosynthetic reaction center contains two tyrosines with different roles in catalysis. The tyrosines, YZ and YD, exhibit differences in kinetics and midpoint potentials, and these differences must be due to noncovalent interactions with the protein environment. Here, studies of YD and YZ and relevant model compounds are described.

Redox-active tyrosine residues have been proposed to be mechanistically important in several enzymes, including ribonucleotide reductase (RNR) [1], photosystem II (PSII) [2, 3], prostaglandin H synthase [4], galactose oxidase [5], glyoxyl oxidase [6], and Mycobacterium tuberculosis catalase-peroxidase [7]. Redox-active tyrosine residues may mediate long range electron transfer (ET) reactions {reviewed in [8]}. Participation in proton transfer (PT) is possible because the oxidation of a protonated tyrosine at neutral pH values is coupled with the deprotonation of the phenolic oxygen [9]. Changes in the pK of the proton-accepting group and in the mechanism of proton/electron transfer can then alter the rate of the reaction [10-13]. In direct coupling reactions, the proton and electron may move simultaneously, sequentially, or their movement may be non-synchronous [14]. In this article, the term proton coupled electron transfer, PCET, will be used to refer to any reaction in which both an electron and a proton are transferred, regardless of mechanism [15, 16].

PSII

PSII contains two redox active tyrosines, YD and YZ, with different functional roles. PSII is the thylakoid membrane protein complex, which carries out the light-catalyzed oxidation of H₂O and reduction of plastoquinone [17, 18]. PSII is found in plants, cyanobacteria, and algae. Electron transfer is initiated when light absorption and energy transfer in the pigment antenna lead to electronic excitation of the primary chlorophyll (chl) donor. Subsequent electron transfer reactions result in the production of a chl cation radical, P₆₈₀⁺, and the sequential reduction of a pheophytin, a plastoquinone, Q_A (a single electron-acceptor), and a second plastoquinone, Q_B (a two electron-two proton acceptor). P₆₈₀⁺ is very unstable and readily generates other oxidized species [19]. One of these oxidized species, YZ^• [2, 3], is an electron transfer intermediate between P₆₈₀⁺ and the catalytic site for water oxidation, the oxygen-evolving complex (OEC) [3, 20, 21].

PSII structural models, derived from X-ray diffraction on three-dimensional crystals, were presented at 3.8-2.9 Å resolution [22-27]. The issue of X-ray-induced Mn reduction has been raised as a potential concern in the interpretation of PSII electron density [25, 28, 29]. PSII is composed of multiple polypeptide components. The CP47 and CP43 subunits are intrinsic, chlorophyll binding proteins [30]. The Mn-stabilizing protein (MSP, psbO, or the 33 kDa protein) is an extrinsic subunit that modulates the properties of the catalytic site [31-33]. Cyanobacterial and plant PSII contain cytochrome b559 [34], as well as other small intrinsic and extrinsic subunits [35]. Two of the hydrophobic PSII subunits, called D1 and D2, bind most of the prosthetic groups that are involved in electron transfer [22-27, 36].

The OEC contains both calcium and manganese [22-27]. Chloride also plays an important role in oxygen evolution {reviewed in [37]} and has recently been identified 6.5 Å from the manganese cluster [27]. The OEC cycles among five oxidation states, called the S_n states, where n refers to the number of oxidizing equivalents stored at the active site [38]. Each S state advancement is driven by one light-driven charge separation in the reaction center ({for examples, see models in [39, 40]}. Oxygen release occurs during the S₃ to S₀ transition from an unstable intermediate, known as the S₄ state. Four flashes are required to produce molecular oxygen from water, and the S₁ state is the dark-adapted state [38]. Therefore, the oxygen-evolving reactions exhibit period four oscillations, with oxygen produced on the third flash and then on every fourth flash [38]. The S state transitions occur in the microsecond to millisecond time regime, with a rate that slows as oxidizing equivalents are stored in the OEC [20, 41].

Two redox active tyrosines, YZ and YD, in PSII

PSII is a prototypical system for the study of redox active amino acids, because two redox active tyrosine residues are involved in light-induced electron transfer reactions, but with different functional roles [19, 42, 43]. Site-directed mutagenesis showed that YZ is tyrosine 161 of the D1 polypeptide and that YD is tyrosine 160 of the D2 polypeptide [44-48]. Both redox active tyrosines are neutral radicals in the oxidized form, suggesting that PCET occurs [2, 3, 49]. YZ mediates electron transfer events between the OEC and the chl donor and is required for oxygen evolution [20, 21, 46, 47]. The other redox active tyrosine, YD, is also oxidized via P₆₈₀⁺ and is in slow redox equilibrium with the OEC [2, 50-52]. YD may be involved in assembly of the OEC [53], but is not required for oxygen evolution [44, 45].

The PSII redox active tyrosines are in C₂ symmetrically related positions in the D1 and D2 subunits (Figure 1). However, only one of the two side chains, YZ, is involved in the electron transfer events that lead to water oxidation. Such structural symmetry, but functional asymmetry, is also observed in the purple bacterial reaction center, which does not oxidize water. In this enzyme, there are two bacteriopheophytin acceptors, positioned with approximately C₂ symmetry, but only one is used in functionally significant electron-transfer events [54].

Figure 1.

The environments of YZ (A) and YD (B) from the 2.9 Å X-ray structure of PSII [27]. In (A), the manganese ions are gray, and the calcium ion is orange.

Functional differences between YZ and YD

The tyrosyl radicals in PSII are distinguished by their different reduction kinetics and midpoint potentials {reviewed in [19, 42, 43]}. YZ and YD had midpoint potentials that differ by ~200 mV [47, 55]. YD^• was stable after illumination for minutes to hours, depending on the PSII preparation, and its EPR signal (Figure 2) was observed as a dark stable signal {for examples, see [2, 43, 56]}. In oxygen-evolving samples, YZ^• was reduced by the OEC on the microsecond time scale, with a rate that depends on S state [20]. In samples from which manganese, calcium, and chloride were removed (OEC-depleted), the EPR lifetime of YZ^• increased to the hundreds of millisecond time scale [57, 58]. EPR signals from YZ^• were detected in OEC-depleted preparations [59] (Figure 2) and in oxygen-evolving PSII [20, 60, 61]. Recent experiments were performed at temperatures >100 K and at high microwave powers, and the YZ^• radical was shown to decay on the millisecond time scale after flash excitation, due to recombination with the PSII acceptor side [60, 61]. At these temperatures, the expected electron spin-spin interaction between the manganese cluster and YZ^• collapsed, due to an increase in the relaxation rate of the metal cluster. It was proposed that YZ^• may act as a hydrogen atom (H^•) abstractor during the S state cycle [62], but in the current X-ray structures, YZ is too far from the OEC to directly abstract hydrogen from substrate water [22-27]. Currently, it is hypothesized that YZ is an intermediary in electron transfer on each S state transition [20, 21].

Figure 2.

Structures and spectra of tyrosyl radicals. (A) shows the dihedral angle at the methylene carbon, with the angles between the hydrogens and the p_Z orbital defined as Θ₁ and Θ₂; (B) model of tyrosyl radical; (C) EPR spectra of the light induced signal from YZ^• and YD^• (or Z and D, solid line) and the dark stable YD^• (or D, dashed line).

EPR studies of isotopically labeled tyrosinate model compounds revealed that tyrosine oxidation occurs from the aromatic ring (Figure 2). Oxidation generated a neutral radical with spin density located on the 1′, 3′, and 5′ carbon atoms and the phenolic oxygen [49, 63]. Rotation around the C_1′-C_β bond (Figure 2) altered the EPR line shape. The EPR lineshapes and g values of YZ^• and YD^• resembled the tyrosyl radical produced in tyrosinate powders at 77 K. The g values of the YZ^• and YD^• were consistent with the deprotonation of both residues (Figure 2) when oxidized [2, 3]. There was one spin of YD^• per reaction center, and up to one spin of YZ^• was generated in OEC-depleted preparations {reviewed in [19, 42, 43]}.

Because YD and YZ have different functions and midpoint potentials, each tyrosine must have a different interaction with its environment. Spectroscopic differences between the tyrosyl radicals were reported [3, 64-68]. Crystal structures of cyanobacterial PSII [24-27] showed that the environments of YZ and YD differ in the detailed placement of the calcium-manganese cluster and hydrophilic amino acids (Figure 1). YZ was 5 Å from the calcium in the OEC, while YD is 28 Å away from the metal cluster. YZ was predicted to hydrogen bond with H190D1 and is 5 Å from D170D1 and 8 Å from R357CP43. YZ may also interact electrostatically with a second histidine (H332D1) residue and a second aspartic acid (D342D1) residue, which coordinate manganese. YD was predicted to form a hydrogen bond with H189D2 and is 7 Å from an D333D2-R180D2 salt bridge. In addition, YD had potential pi-cation interactions with R272 of the CP47 subunit, at 8 Å, and with R294 of the D2 subunit, at 6.5 Å. The corresponding arginines are not found in the environment of YZ. Overall, the pockets are distinguished by increased hydrophobicity near YD.

Mechanism of PCET reactions

The mechanism of PCET may distinguish YD and YZ. Supporting this idea, differential pH effects on the oxidation kinetics of YD and YZ were observed [52]. Marcus electron transfer theory states that the electron transfer rate constant depends on the Gibbs free energy for the reaction, ΔG^o, and on λ, the energy required to reorganize the nuclear configuration from that of the reactants to products [69]. The treatment is based on activated complex theory and simple harmonic potentials. The key parameters can be summarized in a plot of free energy versus reaction coordinate. Two overlapping parabolas represent potential energy surfaces for the reactants in the initial state and products in the final, electron transferred state. Changes in driving force, ΔG^o (the free energy difference between reactants and products) cause vertical displacements of the parabolas; changes in λ cause horizontal displacements. Larger horizontal displacements (larger values of λ) produce larger activation barriers. At constant λ, increases in driving force increase and then decrease the rate, generating the well known inverted region. The maximum rate occurs when the absolute value of the driving force equals λ. The rate of electron transfer varied exponentially with distance, with an exponential factor, β, which is approximately 1.4 Å⁻¹ [70]. However, the protein environment can alter the value of β, the reorganization energy and the driving force to fine-tune the reaction rate [71]. Theoretical treatments for PCET reactions, which are constrained to occur over short distances, have emerged {see for example [72, 73]. A Marcus-type treatment has also been applied to explain PCET [10, 74].

Three pathways for a PCET reaction, involving a redox active tyrosine, can be imagined (Figure 3) [10, 75, 76]. In the first pathway, proton transfer occurs first, followed by electron transfer (PTET). In a second pathway, electron transfer precedes proton transfer (ETPT). In a third pathway, electron and proton transfer occur simultaneously in a concerted manner (CPET). For the first and second pathways, either electron or proton transfer can be rate-limiting. CPET has the advantage of avoiding the production of high energy intermediates, such as the tyrosyl cation radical produced in ETPT.

Figure 3.

Three possible PCET reactions for tyrosyl radical reduction, with an imidazole as the proton donor. Reprinted with permission from [122]. Copyright year 2007 American Chemical Society.

These three mechanisms can be distinguished by the magnitude of the solvent isotope effect {reviewed in [10]}. ETPT shows only a small (<1.3) kinetic isotope effect, while a more significant kinetic isotope effect (1.6-81.0) is expected for CPET [10, 77-89]. PTET will give only a small equilibrium isotope effect (~0.5 ΔpK) in ²H₂O buffers [10, 90, 91].

The pH dependence of the rate constant also helps to distinguish PCET pathways. PTET predicts a pH dependent rate with a rate constant that changes by a factor of 10 per pH unit [10, 89, 92, 93]. ETPT will exhibit no significant pH dependence, if electron transfer is the rate limiting step [89]. Under some conditions, CPET reactions have been reported to be pH dependent [75, 89, 91, 94], although the origin of this pH dependence is controversial {reviewed in [95]}.

Mechanism can also be investigated using temperature dependent studies. These experiments establish the reorganization energy for the PCET reaction, assuming an adiabatic Marcus-type treatment [10, 74]. Substitution of modified electron transfer cofactors with altered potentials and pK is helpful in deducing mechanism. Examples are the use of 3-fluorotyrosine to investigate YZ electron transfer and the use of altered quinones to investigate electron transfer in the bacterial reaction center (see discussions below).

Models for PCET reactions

For tyrosyl radicals, model systems were used to investigate the effect of specific intramolecular interactions in hydrogen bonded, modified phenols [10]. Studies involving phenol derivatives with pendant bases showed that hydrogen bonding decreases the reduction potential of tyrosine [10]. Tyrosine and tryptophan residues were linked to ruthenium photosensitizers [75, 91, 96]. Studies of tyrosine appended to rhenium complexes [76] investigated the pH and buffer dependence of the PCET reactions [91]. Electrochemistry and other techniques have been used to examine PCET reactions for the oxidation of phenols in water [89, 97].

A CPET, rather than the stepwise PTET or ETPT, mechanism was inferred for tyrosine/phenol model compounds under some conditions [10, 75, 76, 91, 97, 98]. For example, in the work of [10, 99], PCET in phenols with pendant, hydrogen bonded bases was studied. The bases were an amine, an imidazole, and a pyridine. The phenol was oxidized with an accompanying intramolecular proton transfer to the hydrogen bonded base. From studies of the dependence of the rate on driving force and temperature, it was concluded that adiabatic Marcus theory can be applied to the PCET reaction. Further, evaluation of the kinetic data led to the suggestion that proton transfer occurs by a CPET mechanism. It was pointed out that the reactions proceed with a large reorganization energy (~2.4 eV or 56 kcal/mole, for one example). The suggested criteria for CPET were threefold. First, the magnitudes of the isotope effects (1.6-2.8) were considered to be consistent with CPET, but not with an ETPT or PTET mechanism. Second, it was concluded that the intermediates produced in ETPT and PTET (the phenoxyl cation radical and the phenolate species) were too high in energy to be accessible during the reactions. Third, the activation energy was observed to depend on the driving force for the reaction [10, 99]. For the reaction rates measured, this was judged to be consistent with CPET, but inconsistent with ETPT and PTET.

A CPET mechanism was also proposed for linked tyrosine-histidine-ruthenium model compounds [75, 84, 100]. For example, in the model compound experiments of [75], photoexcitation of ruthenium reduces an exogenous acceptor. The tyrosine then transfers an electron to the oxidized ruthenium. The experiment was conducted by monitoring the ground state recovery of Ru(II) at 450 nm. For pH values below the pK of the tyrosine (~10), the rate constant increased with pH and had a solvent isotope effect of ~2.0 [75, 82]. Under these conditions, a large reorganization energy, 2 eV, was deduced from studies of the temperature dependence. For pH values above the pK of the tyrosine, the tyrosine was deprotonated, and the rate increased and became pH independent. The reorganization energy decreased to 0.9 eV. The interpretation of these results was that mechanism is CPET below pH 10, but is simple ET, from the unprotonated tyrosine, above pH 10 [75]. It was suggested that a large reorganization energy is a characteristic of the CPET transfer, and that the observed pH dependence arises from the entropy term in the driving force {but see discussion in [95]}.

Calculations were conducted on a model of this system using multistate continuum theory [84]. The solute was described by a multistate valence bond model, the transferring hydrogen nucleus was treated quantum mechanically, and the solvent was represented as a dielectric continuum. The free energy surfaces for ET as a function of a single solvent coordinate or for PCET as a function of two solvent coordinates, corresponding to PT and ET, were calculated. Rate expressions for the ET and CPET reactions were also provided. The pH dependence of the CPET reaction was attributed to the decrease in driving force with pH {[84], but see [89] and [95] references therein). The larger rate for the pure ET reaction was attributed to a smaller solvent reorganization energy for ET and a vibrational overlap factor in the CPET rate expression. The predicted temperature dependence, the solvent isotope effects, and the solvent reorganization energy were reproduced in the calculation [84]. The extension of this approach to more complex systems, which model the PSII reaction center, would be welcome.

Protein and peptide models for PCET

The complexity of enzyme structures, such as PSII, can make it difficult to isolate and manipulate specific variables that influence rate and mechanism. Therefore, the development of simpler peptide/protein electron transfer model systems of defined structure is of value. In one previous approach, small electron transfer proteins such as cytochrome c or azurin, were modified to contain a bound ruthenium ion, with which intermolecular or intramolecular electron transfer was photoinitiated {see, for example, [13, 101] and references therein}.

Another approach is to use a designed, de novo sequence as a framework to investigate electron transfer. The effect of noncovalent interactions on the potential of redox active cofactors, such as hemes, iron sulfur clusters, and other metal centers, has been systematically explored in these peptide models or maquettes {for examples, see refs [102-108]}. For redox active tyrosines, previous work has reported a shift in tyrosine reduction potential in designed helical proteins that contain tyrosine residues [109, 110]. The redox potential of tyrosine was modulated by placement in a non-polar environment and by shielding from proton acceptors [109, 110]. Cyclodecapeptides have also been used to model redox active tyrosine residues in proteins [111].

A beta hairpin maquette exhibits a PCET reaction

Another peptide system in which specific intermolecular and covalent interactions can be probed was recently described. Peptide A consisted of 18 amino acids, IMDRYRVRNGDRIHIRLR, and folded into a beta hairpin [112] (Figure 4). For previous examples of designed beta hairpin peptides, see refs [113, 114]. Peptide A contained a single tyrosine residue (Y5), which was oxidized by UV photolysis or by electrochemical techniques. The experimental results suggested that this peptide exhibits an interstrand PCET reaction between a histidine (H14) and Y5, which are involved in a pi-pi stacking interaction. Substitutions at the histidine were observed to eliminate inflection points in the Pourbaix diagram, which plots peak potential versus pH [112, 115]. This result implied that the inflection points are due to oxidation-induced pK shifts in the histidine. Fits to the data gave pK values for the histidine in the oxidized and reduced forms of the peptide [115]. The extracted pK values suggested a shift from histidine pK 6 in the reduced form of the peptide to histidine pK 9 in the oxidized form. This analysis implied that oxidation of the tyrosine is accompanied by protonation of the histidine in the mid pH region. Although histidine and tyrosine are not directly hydrogen bonded, proton transfer may occur through a bridge of one or more hydrogen bonded water molecules [91, 97, 112, 116]. This peptide exhibits an example of a PCET reaction in which the electron and proton acceptors are two distinct species. Electrochemical data also provided evidence that histidine protonation decreases tyrosine midpoint potential. This effect was attributed to electrostatic stabilization of electron density on the tyrosyl radical phenolic oxygen [112].

Figure 4.

Structure of a beta hairpin peptide, IMDRYRVRNGDRIHIRLR, in which a tyrosine-histidine pi-pi interaction lowers the tyrosine redox potential and a PCET reaction occurs between tyrosine and histidine. (A) shows the primary sequence and some of the predicted hydrogen bonds and cross strand interactions; (B) is the overlap of the 20 lowest energy 2-D NMR structures, with only five amino acid side chains shown (see part A, green); and (C) is the redox active tyrosine and its immediate environment in the averaged, minimized NMR structure. Predicted hydrogen bonds are shown with dashed lines. Reprinted with permission from [112]. Copyright year 2007 American Chemical Society.

YD has a lower midpoint potential than YZ [47, 55], and its protein environment is distinguished by potential pi-cation interactions with arginine residues. In peptide A, the single tyrosine is hydrogen bonded to an arginine residue, R16, and a second arginine, R12, has a pi-cation interaction with Y5. Beta hairpin maquettes were used to evaluate the effect of these hydrogen bonding and electrostatic interactions on the PCET reactions [115]. Optical titrations showed that R16 substitution changes the pK of Y5. Removal of R16 or R12 increased the midpoint potential for tyrosine oxidation. The effects of R12 substitution were consistent with the midpoint potential difference, which is observed for the PSII redox active tyrosine residues. These results demonstrated that a pi-cation interaction, hydrogen bonding, and changes in the nature of PCET reactions can alter redox active tyrosine function. Each interaction may contribute equally to the control of midpoint potential [115].

PCET and YD in PSII

In PSII structures (Figure 1), YD was predicted to hydrogen bond to H189D2 [24-27]. Spectroscopic evidence suggested that proton transfer occurs from a protonated YD to the nearby histidine, H189D2 [117], and that the neutral radical, YD^•, is hydrogen bonded to this histidine [118]. In site directed mutants at H189D2, reconstitution of imidazole accelerated the rate of YD^• reduction. This result implied that YD proton transfer is rate limiting and that proton and electron transfer are kinetically coupled [117]. Mutations at H189D2 substantially changed the EPR lineshape of YD [119-121]. At pH 7.5, reaction-induced FT-IR spectroscopy showed a perturbation of the H189D2 NH stretching vibrations. The perturbation was attributed to a proton transfer reaction from the protonated imidazole side chain of H189D2 to YD^• [117].

The effect of solvent isotope exchange on the reduction kinetics of YD^• was examined. Experiments were conducted as a function of pL (the negative logarithm of the lyonium ion concentration). The kinetic isotope effect (KIE) on YD^• reduction was measured by EPR spectroscopy [122]. YD^• was detected independently of YZ^•, because YZ^• decayed in an interval that was not used in kinetic analysis. The rate of YD^• reduction was slow, and so it was assumed that PT, which occurs over short distances {discussed in [10] and references therein}, does not limit the reaction rate. These studies showed that YD proton transfer reactions proceed with different mechanisms at high and low pH [122, 123]. YD^• reduction exhibited a pL-dependent rate constant, with a minimum at ~pL 7.0-7.5 (Figure 5). At high pL (>7.5), the values of the KIE were greater than 2.0 and were consistent with a CPET mechanism. At low pL, smaller KIE’s were observed, and this result was attributed to a PTET mechanism in which the protonation of YD^• occurs first. The rate acceleration observed at low pH was consistent with PTET.

Figure 5.

pL dependence of the YD^• reduction rate, as determined by EPR spectroscopy, in ¹H₂O (black) and ²H₂O (blue) buffers. This slow phase corresponds to ≥85% of the signal amplitude. Reprinted with permission from [122]. Copyright year 2007 American Chemical Society.

A proton inventory was recorded to determine the number of protons transferred at high pH [123]. This technique uses changes in the mole fraction of ²H₂O to determine the number of protons transferred in the rate-limiting step of a reaction [124]. The decay of YD^• was monitored as a function of ²H₂O mole fraction at pL 8.0, where there is a significant isotope effect. Again, EPR transients were recorded on a time scale in which YZ^• does not contribute to the data. When the rate was plotted versus ²H₂O content, hypercurvature was observed. This hypercurvature provided evidence for the existence of multiple, proton-donation pathways to YD^• (Figure 6). For comparison, a rocking model [125] in which histidine donates and then reaccepts a proton would give a linear proton inventory, corresponding to the transfer of one proton in the transition state. Simulation of the proton inventory data suggested that at least one of the proton transfer pathways involves the transfer of more than one proton (Figure 6). Modeling of the data supported the conclusion that there are two competing proton transfer pathways, one involving one proton and the other a multiproton (≥3 proton) pathway (Figure 6). It was proposed that one pathway involves H189D2 as proton donor and that the other pathway involves water as the proton donor [123].

Figure 6.

Two possible models for proton transfer involving YD^•, based on the observation of hypercurvature in the proton inventory. The arrows indicate proton, not electron, movement. A. In pathway 1, histidine (blue) is involved in the multiproton pathway, which must involve three or more protons, of which two are shown. In pathway 2, a water molecule (red) is proposed to act as a single proton donor. B. In pathway 1, histidine (blue) is proposed to act as a single proton donor. In pathway 2, a chain of water molecules (red) are involved in the multiproton pathway, which must involve three or more protons, of which only two are shown. Reprinted with permission from [123]. Copyright year 2009 American Chemical Society.

H190D1 and YZ in PSII

In PSII structures, YZ was predicted to hydrogen bond to H190D1 [24-27]. YZ was suggested to be protonated at high temperatures [61] and will therefore lose a proton when oxidized. From chemical complementation studies, it was proposed that H190D1 is the proton acceptor for YZ and that the hydroxyl proton of YZ^• remains bound to H190D1 during the lifetime of the radical [126, 127]. In these studies, the rate of YZ oxidation was monitored by optical studies of P₆₈₀⁺ reduction at 432.5 and at 810 nm. The rate of YZ^• reduction was monitored by optical studies at 242 nm. Mutants at H190D1 did not bind the OEC and are therefore OEC-depleted [126, 128, 129]. The rates of YZ oxidation and reduction slowed in site directed mutations at H190D1, but accelerated when imidazole and other small bases were reconstituted. An imidazole effect was observed both in wildtype and mutant PSII [126]. The mutant result suggested that YZ proton transfer is rate limiting and that electron and proton transfer are coupled for the oxidation and the reduction reaction. The fact that imidazole addition gave increased rates both for YZ oxidation and reduction even in wildtype PSII, which contains H190D1, was attributed to a suboptimal interaction between YZ and H190D1, in the absence of the manganese cluster [126, 127]. High concentrations of imidazole also accelerated P₆₈₀⁺ reduction in the Y161D1F mutant, which lacks tyrosine Z. The etiology of this effect is unclear.

Contrasting kinetic results were obtained by room and low temperature EPR studies in a H190D1Q mutant [129]. In that work, the EPR lineshape of YZ^• was only slightly altered by the mutation {see also [130]}. This was in contrast to the situation in the H189D2 mutant, in which substantial YD^• lineshape changes were observed (see above). The reason for this difference between the D1 and D2 mutants is not yet clear. At −10°C, , the yield of YZ^• in His190D1Q was found to be lower than that of wildtype, but only by a factor of 1.6, and the reduction kinetics were not significantly altered by the mutation. At room temperature, the YZ^• yield was decreased by a factor of 2.6 in this mutant, but again, the kinetics of YZ^• reduction appeared similar to wildtype [129]. These results suggested that an alternant proton donor, other than H190, is active in YZ^• reduction reactions under these conditions. One possible explanation is that the identity of the proton donor/acceptor is different for the oxidation and reduction reaction [129]. This could occur if YZ^• oxidation induces a local structural change, which changes its proximity to local bases and acids.

If oxidation of YZ triggers a conformational change, temperature dependence might be expected in the redox reactions. Significant temperature dependence was observed in the yield of the YZ^• EPR signal in the His190D1Q mutant [129]. The yield at 4 °C was 70% of the −10 °C yield, and the yield at 21 °C was 40% of the −10 °C yield. However, there was no significant change in the overall kinetics of YZ^• decay at the three temperatures. This observed temperature dependence in the His190D1Q mutant was a reversible phenomenon, with a recovery of 80% of the yield when the 21°C sample was reequilibrated at −10°C. The field-swept EPR spectra acquired under continuous illumination were indistinguishable at all three temperatures (data not shown). In summary, EPR analysis of the H190D1Q mutant suggests complexity in the YZ proton transfer reactions, which is congruent with some of the interpretations described below.

PCET reactions and YZ

Based on EPR studies, it was suggested that the mechanism of YZ PCET is different in oxygen-evolving and OEC-depleted PSII [61]. OEC removal may cause changes in the environment of YZ^•, including more disordered hydrogen bonding to the tyrosyl radical [61, 65, 67, 68, 131]. Also, in the presence of the OEC, YZ^• is in a more electropositive environment, compared to YZ^• in an OEC-depleted preparations [61]. A synopsis of EPR and optical studies on PCET reactions in oxygen-evolving and OEC-depleted PSII is given below. Investigations have focused both on the oxidation and reduction reaction.

EPR spectroscopy investigates solvent isotope effects in OEC-depleted and oxygen-evolving PSII

Solvent isotope and pH effects on YZ redox reactions were reported using EPR spectroscopy to study spinach PSII. In [132], an EPR study was reported on oxygen-evolving and OEC-depleted PSII at room temperature. The kinetics of YZ^• reduction were monitored. No significant solvent isotope effect was observed on YZ^• reduction at pL 6.0 in OEC-depleted PSII, but KIE values of 2.9, 1.3, and 1.6 were reported for the S₁ to YZ^•, S₂ to YZ^•, and S₃ to YZ^• reduction reactions. These isotope effects could be due to PCET events at the OEC or at YZ. The value for the S₁ to YZ^• reduction reaction was larger than the KIE measured by optical spectroscopy, which gave values of 1.3-1.4 for each S state transition in the same type of PSII preparation [133]. This discrepancy could be caused by a difference in the extent of solvent isotope exchange in the two cases. In [134], another room temperature EPR study of YZ^• reduction from the OEC was reported. In this case, PSII retained the manganese cluster, but Sr⁺² was substituted for Ca⁺² in order to slow the rate of YZ^• reduction. A KIE of 1.4 was reported at pL 6.5, suggesting that metal ion substitution did not change the mechanism.

An EPR study reported a proton inventory for the YZ oxidation reaction in OEC-depleted spinach PSII [135]. The isotope and pH effects were assessed indirectly by measurement of YZ^• yield under illumination. The findings were that one proton transferred during the rate-limiting step of the reaction and that the oxidation rate of YZ was pH dependent. The derived values were reported as KIE=1.4 at pL≤5 and pL>7 and as KIE=5 at pL 6.0. Thus, this experiment supports the conclusion that there is a small isotope effect on the YZ oxidation reaction. A more detailed interpretation of the mechanism awaits further studies.

Optical spectroscopy suggests two different proton acceptors for YZ in OEC-depleted PSII and shows no significant solvent isotope effect for YZ oxidation in oxygen-evolving PSII

Optical spectroscopy was used to study YZ oxidation in oxygen-evolving and OEC-depleted PSII from pea [136]. The reduction of P₆₈₀⁺ was monitored. In OEC-depleted PSII, the overall decay time was pH dependent and slowed as the pH decreased. The data were fit with three exponentials. The two fastest phases were attributed to electron transfer between YZ and P₆₈₀⁺. The fast phase (1 microsecond half decay time) was largely pH independent, and the slower kinetic component (35 microsecond half decay time at pH 4) exhibited only a five-fold decrease in rate from pH 8.0 to pH 4.0. However, the amplitudes were pH dependent, with the fast phase dominating at high pH. The fast phase was insensitive to solvent isotope exchange; the slower phase showed an isotope effect of 2.5. In oxygen-evolving centers, the rate of YZ to P₆₈₀⁺ electron transfer occurred in the nanosecond time regime and showed no significant isotope effect. These results concerning the nanosecond rates agree with previous reports {see [133, 137, 138] and references therein}. In oxygen-evolving centers below pH 5, the nanosecond components were replaced by slower components, attributed to a pH-induced inactivation of PSII centers {[136], but see [138]}.

The experiments of ref [136] on OEC-depleted pea PSII were reinterpreted in [75], based on comparison to new model studies. Similarities in reorganization energy and in the pH dependence of the rates were observed when OEC-depleted PSII was compared to ruthenium-tyrosine containing compounds. In the model compounds, it was concluded that a CPET mechanism applies at low pH and that at pH values above the pK of tyrosine, pure ET occurs. Comparison to data from OEC-depleted PSII showed a similarity in pH dependence and activation energy in the low pH regime. This similarity led to the suggestion that a CPET mechanism applies at pH <7.6 and a pure ET mechanism applies at pH >7.6 in PSII. At pH<7.6, proton release was proposed to occur directly into bulk water. At pH>7.6, it was suggested that a hydrogen bond forms between H190 and YZ; this hydrogen bond was suggested to facilitate electron transfer by decreasing the reorganization energy {see also [84]}. The hydrogen bond between H190 and YZ was suggested to be broken at lower pH [75]. Thus, this interpretation suggests two different proton acceptors for YZ in OEC-depleted PSII. The nature of proton transfer in oxygen-evolving PSII is not clear, because no significant isotope effect was observed. More experiments are necessary to elucidate this question.

Optical spectroscopy on cyanobacterial PSII shows significant solvent isotope effects and pH dependence for YZ redox reactions

The cyanobacterial Y160D2F mutant lacks YD, and thus the YZ redox reactions can be studied in a background that lacks the stable tyrosyl radical signal from YD. However, the mutation may alter proton transfer pathways on the donor side of PSII [139]. In OEC-depleted PSII from the Y160D2F mutant, the oxidation of YZ was monitored by decreases in the P₆₈₀⁺ signal [131]. The KIE value was 3.6 at pL 7.0 (maximal) and was pH dependent. The oxidation rate also depended on pL, with the reaction rate increasing at higher pL values. The rate constants in ¹H₂O varied from 30.6 ms⁻¹ at pH 5.5 to 1260 ms⁻¹ at pH 9.5. There results were interpreted with a mechanism, which is electron transfer limited at high pH values and conformationally gated. 0at low pH values. However, results from fluorotyrosine labeling (described below) suggest a reinterpretation of these results. The rate of YZ reduction was also measured, by monitoring its recombination with Q_A⁻ at 325 nm. The KIE for the reduction reaction was 2.0 and was smaller than the KIE for the oxidation reaction. The rate of the reduction reaction showed less dependence on pL. It was concluded that there was no rate limitation on proton transfer in the YZ reduction reaction [131]. However, in this case, the origin of the isotope effect is unclear.

In a study of OEC-depleted PSII from wildtype cyanobacteria (retaining YD), the rate of YZ oxidation and reduction were also monitored by optical spectroscopy. The kinetics of P₆₈₀⁺ decay, as monitored at 810 nm, were fit with four exponentials. Again, pH dependence was observed. While the individual rate constants did not vary extensively with pH, the relative amplitudes showed variation [127]. At pH 7.5, the k⁻¹ values were 340 ns, 3.4 μs, 21 μs, and 120 μs. Details of the observed pH dependence were different from the measurements of [131]. The isotope effects were also smaller and pH independent (1.7-2.6). Differences between ref [127] and [131] may be due to the use of the YF160D2 mutant in [131]. However, ref [127] attributed the difference to lower time resolution in the measurements reported in [131]. In ref [127], addition of imidazole and other bases accelerated the rate of YZ oxidation and reduction, even in these wildtype PSII preparations. The experiments were interpreted as supporting either a CPET mechanism or a mechanism in which proton transfer is rate limiting [127]. The observed influence of exogenous bases on electron transfer in wildtype PSII is interesting. The explanation was that when manganese is removed, the hydrogen bond between YZ and H190 is broken. The suggestion was that the rate acceleration in wildtype is due to access of exogenous proton donors to YZ^•.

To summarize, studies of mutant and wildtype cyanobacterial PSII agree that the rate of YZ oxidation accelerates with increasing pH. Isotope effects have been reported on the oxidation and reduction reactions [127, 131]. Small differences in rates and isotope effects, when pea [136] and cyanobacteria [127, 131] were compared, may reflect biochemical differences between the preparations. Alterations in the PCET pathway may complicate interpretation of studies on the Y160D2 mutant [131].

Fluorinated tyrosine labeling suggests a YZ oxidation mechanism in OEC-depleted PSII

A study of PCET has been reported for YZ using fluorinated tyrosine [140]. Fluorinated tyrosine has been used previously to investigate the mechanism of PCET in RNR [141, 142]. In PSII, the extent of fluorination was reported as 75%, based on analysis of the altered electron spin echo spectrum. Fluorinated tyrosine has both an altered pK and midpoint potential [141]; therefore, global incorporation of fluorinated tyrosine is expected to change the pK and midpoint potential of YZ. In some PSII experiments, a site directed mutant was employed in which a histidine mutation changes the midpoint potential of P₆₈₀⁺. The reduction of P₆₈₀⁺ was monitored by optical spectroscopy, and the samples were OEC-depleted cyanobacterial PSII. The pH dependence of the rate was not altered in the fluorotyrosine-containing samples. Because the pK should decrease by 1.5 units in the fluorotyrosine derivative, it was concluded that the pH dependence reflects a base other than YZ. The rate at pH 9.2 was observed to increase in the fluorotyrosine samples, consistent with the pK change altering a rate-limiting step in the reaction. The activation energy, as assessed by the temperature dependence of the rate, decreased by 110 meV, which corresponds to the expected change in ΔG for the proton transfer reaction in fluorotyrosine. This was distinct from observations with the histidine mutant, which was assumed to change only the driving force for the reaction. From this, it was concluded that the change in activation energy is due to the pK change in fluorotyrosine, not from a difference in driving force. To explain these data, a CPET mechanism was proposed for YZ at low pH, and a sequential PTET mechanism was proposed at high pH [140]. It was concluded that the mechanism of PTET is limited by a proton transfer reaction. To explain the pH dependence, the authors proposed that the network of hydrogen bonds around YZ changes as a function of pH. It was also suggested that an alternate proton acceptor may be available, other than H190, at low pH.

To summarize, at low pH values, both [140] and [75] predict a CPET mechanism for YZ oxidation in OEC-depleted PSII. Fluorotyrosine labeling suggested a PTET mechanism at high pH and complexity in the proton transfer pathway [140]. The effect of OEC removal on YZ PCET remains to be evaluated.

Are redox reactions at YZ coupled with a conformational change?

Reaction-induced FT-IR and EPR spectroscopies were used to study tyrosine oxidation in tyrosinate powders and in tyrosine-peptides [143, 144]. This model compound work suggested that Y^• reduction is accompanied by a conformational change. In model compounds, Y^• can be generated by UV photolysis at liquid nitrogen temperatures (77 K). The difference or reaction-induced FT-IR spectra reflect the changes in CO and ring stretching vibrations, which are associated with formation of the radical. When the reaction-induced FT-IR spectra acquired on tyrosinate and on peptides were compared, the peptide spectra exhibited unique bands in the amide I (1650 cm⁻¹) region of the spectrum [143-145]. The frequencies of the bands were found depend on amino acid composition and/or sequence. Further, an EPR transient, with a lifetime on the seconds time scale, was observed at 85 K in tyrosinate powders [144].

The EPR transient was attributed to a conformational change, which is expected from DFT studies. At room temperature, DFT calculations predict two stable conformers at the beta methylene bond for tyrosinate, but only one stable conformer for the tyrosyl radical [145, 146]. For tyrosine, conformer A had a ring dihedral angle of +168°, and conformer B had a ring dihedral angle of −69°. ESEEM and FT-IR studies suggested that the B conformer is favored by solvent interactions [143, 146]. For tyrosyl radical, DFT calculations predicted one stable conformer, with a ring dihedral angle of +165°. Therefore, when the radical is generated at low temperature, the initial, populated radical states have ring dihedral angles representative of the tyrosine ground state. The initially populated B state will be unstable at higher temperatures [147] and its relaxation could give rise to the EPR transient [144].

What is the origin of the amide I bands in the model Y^• FT-IR spectra? Interestingly, amide I bands have also been detected in FT-IR studies of the tyrosyl radical in RNR. Reaction-induced FT-IR spectroscopy was used to study the hydroxurea-induced reduction of tyrosyl radical, Y122^•, in the beta subunit [148]. EPR spectroscopy had previously provided evidence for a conformational change at the beta methylene position [149]. Reaction-induced FT-IR spectroscopy showed that reduction of the Y122^• radical was accompanied by amide I contributions, which had similar frequencies to those observed in a pentapeptide, RSTYH [148]. RSTYH is the primary sequence surrounding Y122 [144].

One possible origin for the perturbation of the amide bond frequencies could be spin delocalization to the amide nitrogen [150]. However, this was not consistent with subsequent ESEEM studies of model compounds [146]. An alternate origin for the changes in amide I frequencies is an alteration in amide bond ionic character. Rotation around the amide bond changes the electronegativity of the amide nitrogen, the ionic character of the CN bond, and the force constants for the CN stretching modes [151]. Such a change in amide bond planarity could be the conformational change detected and could be driven by the change in electron density, which makes the phenolic oxygen atom more negative in the radical state [112].

In PSII, reaction-induced FT-IR spectra of YZ have also detected amide I frequencies, perturbed by YZ oxidation and reduction [150, 152, 153]. Structural changes preceding the S state transitions have been reported [154, 155]; these changes may correspond to alterations in amino acid protonation state and may be driven by YZ redox or conformational reactions. If oxidation of YZ causes a conformational change, such an alteration could expose the YZ phenolic oxygen to a different acid/base environment. For example, after oxidation of YZ and protonation of H190D1, such a conformational change could rotate YZ^• towards positively charged species, such as Arg357 or the calcium ion (Figure 1). For recent suggestions concerning the role of calcium and Arg357 in the water oxidation cycle see refs [156, 157]. After YZ^• reduction occurs, a decreased distance from a positive charge would stabilize a negative charge on YZ⁻, decreasing its pK. Because Y_Z⁻ is now in a new microenvironment with a potentially different pK, a different species could protonate the tyrosyl radical. Thus, a local conformational change could control PCET reactions involving the redox active tyrosine and could provide a mechanism to translocate protons [156]. If YZ is conformationally flexible, then variations in experimental results could be due to small changes in biochemical preparations, pH, manganese content, or measurement temperatures. The idea that YZ is conformationally flexible in oxygen-evolving PSII has been proposed previously [67].

Do redox reactions at the OEC alter YZ PCET reactions?

As exhibited in the discussion above, YZ PCET reactions are complex, and there is no consensus yet concerning mechanism, at least in oxygen-evolving PSII. Redox-linked changes in the OEC may influence the protonation state of YZ and PCET reactions [61]. A speculative model giving an example of such a redox-linked reaction is shown in Figure 7. PCET to YZ^• is depicted as ETPT, i.e., two sequential steps, in which a proton transfer follows a rate-limiting electron transfer. However, as discussed above, other possible mechanisms cannot be excluded at the present time. In Figure 7, transfer of protons from the oxidized OEC to a nearby base, B₁, modulates the pK of B₂, the immediate proton donor for YZ^•. Redox-linked changes in the pK of B₂ ensure that YZ^• reprotonates and therefore has the high midpoint potential [158] necessary to drive S state advancement {see discussion in [61]}. This idea implies that the microenvironment of YZ and pK of proton donor/s acceptors may be S state dependent. More experiments on oxygen-evolving PSII will be necessary to evaluate YZ PCET reactions.

Figure 7.

Speculative two base model for YZ PCET reactions in the presence of the OEC, adapted from [61]. The reduction and protonation of YZ^• may occur in these two sequential steps (ETPT) as shown, or by a PTET or CPET mechanism. In the first step (YZH oxidation), YZ is oxidized and loses an electron and a proton to form a neutral tyrosyl radical. B₂ is the amino acid side chain that accepts the proton, forming H⁺B₂. In the second step (S state advancement), the OEC is oxidized by YZ^•, generating YZ⁻, and the pK of a second amino acid side chain, B₁, is altered so that B₁ protonates, forming H⁺B₁. In the third step (YZ⁻ protonation), generation of the positive charge on H⁺B₁ changes the pK of H⁺B₂ so that YZ⁻ protonates to form YZH.

Comparison to PCET involving the quinone acceptors

Comparison can be made to PCET involving quinone acceptors in the bacterial reaction center, in which the mechanism of PCET has been elucidated. PCET events on the PSII acceptor side resemble acceptor side transfer in the bacterial reaction center.

In PSII and the bacterial reaction center, light induced electron transfer leads to the sequential reduction of two quinone molecules, Q_A and Q_B. Two sequential photo-induced reductions occur to generate a reduced quinol, Q_BH₂, in the Q_B site. While Q_B is a two electron, two proton acceptor, Q_A is a single electron acceptor and is not protonated {reviewed in [159]}. Unlike PSII, the bacterial reaction center does not split water and lacks the redox active tyrosines. Instead, a cytochrome acts as an electron donor. Substitutions of quinones at the Q_A and Q_B sites in the bacterial reaction center led to the elucidation of PCET reactions between these electron acceptors. The quinone substitutions changed midpoint potential and pK.

For the first electron transfer from Q_A⁻ to Q_B, it was observed that the rate constant did not depend on driving force, although a substantial activation barrier had been deduced for this reaction {see [160] and references therein}. These results suggested that electron transfer occurs alone, with no proton transfer, and that a protein conformation change controls the rate [160]. This conformation change was attributed to an alteration in the position of Q_B, which was observed in crystals under illumination [161]. For the second electron transfer from Q_A⁻ to Q_B⁻, which involves proton transfer, it was concluded that reversible proton transfer occurs, followed by rate limiting electron transfer [92]. Thus, a two step PTET mechanism applies. The second protonation reaction, to form Q_BH₂, follows the production of Q_BH⁻. The semiquinone anion radical, Q_BH⁻, was detected in the Q_B site of reaction centers, which contained a rhodoquinone with an altered pK value [93]. Comparison with donor side electron transfer in PSII suggests that quinone cofactors catalyze step-wise transfers, while YD and YZ involve CPET mechanisms.

Conclusions

Control of catalysis can occur through modulation of PCET reactions. Both the electron and proton tunnel, but the proton transfer is constrained to occur over short distances. When the distance is short, the proton and electron may be transferred together. However, when the distance is long, separate or orthogonal electron and proton acceptors may be used [11]. Modification of the proton transfer reaction can be a key feature in determining the electron transfer rate. In PSII, much remains to be determined about how the responsive protein matrix participates in PCET reactions. Model compounds and model peptides provide an arena, in which hypotheses generated from PSII can be tested. For example, a linked tyrosine-histidine-ruthenium model compounds mimic PCET reactions in OEC-depleted PSII [75, 84, 100]. A beta hairpin peptide [112, 115] exhibits a PCET reaction between tyrosine and histidine and shows a functional difference induced by a pi-cation interaction, which is consistent with expectations from the PSII structure. The mechanism of PCET may help to distinguish YD and YZ, but more information is necessary concerning PCET in oxygen-evolving PSII samples.

In this article, progress in understanding proton coupled electron transfer (PCET) in photosystem II, the photosynthetic water oxidizing complex, is reviewed. PCET may be catalyzed by redox active tyrosines. Photosystem II provides a paradigm for the study of redox active tyrosines and PCET, because this photosynthetic reaction center contains two tyrosines with different roles in catalysis. Here, studies of YD and YZ and relevant model compounds are described.

Abbreviations

ET

electron transfer

KIE

kinetic isotope effect

L

lyonium ion

OEC

oxygen-evolving complex

PCET

proton coupled electron transfer

PSII

photosystem II

PT

proton transfer

RNR

ribonucleotide reductase