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Functional Water Networks in Fully Hydrated Photosystem II

Cite this: J. Am. Chem. Soc. 2022, 144, 48, 22035–22050
Publication Date (Web):November 22, 2022
https://doi.org/10.1021/jacs.2c09121

Copyright © 2022 The Authors. Published by American Chemical Society. This publication is licensed under

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Abstract

Water channels and networks within photosystem II (PSII) of oxygenic photosynthesis are critical for enzyme structure and function. They control substrate delivery to the oxygen-evolving center and mediate proton transfer at both the oxidative and reductive endpoints. Current views on PSII hydration are derived from protein crystallography, but structural information may be compromised by sample dehydration and technical limitations. Here, we simulate the physiological hydration structure of a cyanobacterial PSII model following a thorough hydration procedure and large-scale unconstrained all-atom molecular dynamics enabled by massively parallel simulations. We show that crystallographic models of PSII are moderately to severely dehydrated and that this problem is particularly acute for models derived from X-ray free electron laser (XFEL) serial femtosecond crystallography. We present a fully hydrated representation of cyanobacterial PSII and map all water channels, both static and dynamic, associated with the electron donor and acceptor sides. Among them, we describe a series of transient channels and the attendant conformational gating role of protein components. On the acceptor side, we characterize a channel system that is absent from existing crystallographic models but is likely functionally important for the reduction of the terminal electron acceptor plastoquinone QB. The results of the present work build a foundation for properly (re)evaluating crystallographic models and for eliciting new insights into PSII structure and function.

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1. Introduction

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Water actively determines the internal structure of enzymes, mediates proton transfer, regulates redox properties, and even participates as reactant in chemical transformations. Nowhere is this better exemplified than in photosystem II (PSII), the membrane-embedded pigment–protein complex that harnesses sunlight to drive water oxidation and plastoquinone reduction. (1−10) Every chemical transformation in PSII, from the four-electron water oxidation at the oxygen-evolving complex (OEC) to the two-electron reduction of the mobile electron carrier plastoquinone, is intimately coupled to water management and proton transfer. Thus, atomic-level elucidation of enzymatic functions relies on precise understanding of water channels and networks within PSII. (11−37)
Crystallographic studies provide crucial structural information in this respect. (38−43) To the extreme, apparent alterations of crystallographic water positions in time-resolved studies have even been used as basis for speculations on mechanistic scenarios. (33,44−47) However, the information content of crystallographic models of PSII is constrained by more than medium resolution and inherent technical limitations in locating water molecules or identifying their dynamics. Dehydration is employed in membrane protein crystallography to improve packing and reduce mosaicity, thus facilitating structure determinations and enhancing resolution. (48−55) On the other hand, loss of water affects the reaction center absorption profiles, inhibits the water oxidation cycle at the OEC, hampers electron transfer among plastoquinones, and compromises overall oxygen evolution activity. (56−61) Despite impairment of physiological function, post-crystallization dehydration is not undesirable because it is a technically beneficial procedure that opened the way toward high-resolution structures of PSII. Nevertheless, the dehydrated sample is, by definition, insufficiently representative of the physiological state even if the global integrity of the enzyme is maintained. The consequences can be far-reaching if reduction in water content leads to inaccurate representation of functionally critical regions such as the active sites of water oxidation and plastoquinone reduction.
In this work, we show that all of the above issues arise in available crystallographic models of PSII. To recover missing information on hydration, we construct a membrane-embedded model of PSII (Figure 1) and employ a multistage hydration procedure coupled to unbiased molecular dynamics (MD) simulations on high-performance graphics processing units. This enables us to reconstruct an approximated physiological hydration structure of cyanobacterial PSII. On this basis, we evaluate the hydration state of crystallographic models. We quantify the extent of dehydration in conventional (synchrotron crystallography) and more recent X-ray free electron laser serial femtosecond crystallography (XFEL-SFX) models, and we show that the latter are drastically dehydrated, not only in a global sense but also internally around the OEC and on the acceptor side of the enzyme. Subsequently, we map all water channels associated with the oxidative and reductive endpoints of PSII. Thanks to the employed simulation time scale, we are able to document new water channel systems and entry/exit points, some of them transiently formed, and describe how they are gated by protein conformational changes. Finally, we characterize a channel system in the stromal side of PSII that is absent from crystallographic models but has an obvious functional role, thus establishing the physiological hydration state of the acceptor side and offering a structural basis for discussing the hydration of the non-heme iron site and the mechanism of plastoquinone reduction.

Figure 1

Figure 1. (a) Model of the membrane-embedded PSII monomer. Key protein subunits are labeled. The location of the oxygen-evolving complex and the non-heme iron are indicated. The PsbO, PsbU, and PsbV proteins cap cyanobacterial PSII on the lumenal side of the membrane. (b) Redox-active cofactors in the core proteins of PSII and the two reactions accomplished at the oxidative and reductive termini. The arrows indicate the normal flow of electrons.

2. Results and Discussion

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2.1. Hydration and Molecular Dynamics Simulations

The diffusion time scale of bulk water to internal cavities can be very long and requires extensive sampling of protein conformations. Therefore, conventional MD simulations need to be performed for longer than typical time scales to accumulate information about the dynamics of buried water molecules. The calculations scale unfavorably with the increasing size of the system, especially for the complete membrane-embedded PSII model. Alternative strategies such as the three-dimensional reference interaction site model (3D-RISM) or integration of the Monte Carlo (MC) method with molecular dynamics (for instance, the MC/MD module implemented in AMBER) have shown tremendous capabilities in predicting the hydration content or solvent-accessible sites inside inhomogeneous environments. In this work, we employed the 3D-RISM (62,63) and MC/MD (64) techniques to hydrate internal cavities of PSII prior to production simulations. 3D-RISM is an integral statistical theory of molecular liquids, which can be used to obtain the 3D density distribution of solvent around a solute, while the MC/MD method is based on the integration of the Metropolis Monte Carlo translational movement of water from the bulk to internal cavities during the MD simulation.
The PSII crystal structure by Umena et al. (41) (PDB ID: 3WU2, monomer A) was used as the starting point. We placed emphasis on completing all structural details that are missing from the crystallographic structure in order to make our computational model as representative of the natural system as possible. The prepared PSII structure was embedded in a large patch of lipid bilayer. In addition, a larger than typical dimension of the water box was used along the membrane normal to eliminate any artificial protein–protein interactions during the MD simulations. The setup is fully described in the Supporting Information.
Given that major effort is devoted to properly hydrate the system before initiating production simulations, a thorough equilibration procedure is followed to ensure proper hydration. As a first step, we kept all crystallographically resolved (1506) water molecules. In a second step, we used the 3D-RISM technique coupled with the placevent module (65) to further identify explicit water positions based on the 3D distribution density. A total of 5404 water molecules were predicted using the latter procedure. The majority of waters were found on the protein–bulk interface, but significant water content was also placed in internal protein cavities. Importantly, we found 32 new water positions within 20 Å around the OEC. This single point 3D-RISM calculation was performed on the crystallographic configuration of the PSII protein complex. However, to fully reverse the dehydration of the crystallographic model, it requires a dynamic treatment of hydration on the computational side, that is, rehydrating in conjunction with time evolution of the system. Following a thorough structural relaxation procedure, we thus performed MC/MD calculations to fill or vacate internal cavities based on energetic criteria. Complete algorithmic details are described by Gilson and coworkers. (64) The schematic workflow of the complete hydration procedure is shown in Figure 2a. Thereafter, we conducted MD simulations for 65 ns to equilibrate the complete system, lipid-bilayer, and specifically to allow the newly added water molecules to reach equilibrium with the bulk. Once the system was completely hydrated and fully equilibrated, we initiated 200 ns of production MD simulation for final data acquisition (Figure 2b). Complete technical details on the hydration protocol, equilibration, and production MD simulations are provided in the Supporting Information.

Figure 2

Figure 2. (a) Flow-chart describing the simulation procedure employed in the current work. (b) Time evolution of root mean square deviation (RMSD) of Cα atoms of the PSII monomer during production simulations (data recorded every 2 ps, excluding the highly flexible loops). (c) Time evolution of the hydration content of the PSII monomer. All water molecules in close contact (∼2 Å) with PSII were selected in the counting (data recorded every 50 ps, hydration content around the highly flexible loops not considered). (d) Time evolution of water content within a sphere of radius 20 Å centered around the OEC (data recorded every 2 ps). Corresponding datasets that include the water content associated with the highly flexible loops are provided in the Supporting Information.

The total hydration content of the PSII monomer is depicted in Figure 2c. All water molecules in close contact (∼2 Å) with PSII were selected in the counting. We observe that the total hydration content of the enzyme remains stable during the simulation. The PSII monomer can hold a minimum of 1897 and a maximum of 2087 number of water molecules, which depends on the conformational state of the protein. In addition, we observe that the conformational flexibility of the protein influences the local architecture of water channels. As a demonstration of this point, we chose a spherical region centered around the OEC with a radius of 20 Å, a distance that covers the maximum internal protein volume while completely avoiding the bulk and transmembrane regions. We observe that the time evolution of the hydration content within this volume remains stable (Figure 2d), even though discrete fluctuations are observed. These reflect the dependence of water content on protein flexibility, manifested as conformational gating of water channels by inter- and intra-subunit salt bridges, side chain rotation of amino acids, and the conformational dynamics of C-terminal residues. An atomistic description of such elements is discussed in the following.

2.2. Comparison with Crystallographic Models

Although water is obviously lost by dehydration procedures, (48,49) it is hard to estimate the actual extent of water content reduction. In Table 1, we provide a view into this problem by comparing the number of explicit water molecules resolved in various crystallographic models (33,41−47,66) and two recent cryo-electron microscopy (cryo-EM) structures (67,68) with the results of our MD simulations. This can serve as an indicator for the wetness of the system. Deviation from the MD hydration content is not a strictly quantitative measure of sample dehydration because the difference from the MD reference cannot be uniquely attributed to the real absence of water. Experimental resolution limitations and incomplete correspondence between crystallographic and computational models introduce ill-defined uncertainties. Nevertheless, the comparison does provide a meaningful, if only semi-quantitative, indication of how far the crystallographic models are from a fully hydrated state.
Table 1. Total Number of Water Molecules and Hydration Content Within 20 Å from the Oxygen Evolving Complex (OEC) Derived from the Molecular Dynamics Simulations of the Fully Hydrated PSII Monomer Compared to the Number of Water Molecules Present in Selected Crystallographic Models of PSII from Synchrotron (SX) and Free Electron Laser (XFEL) Sources and from Cryo-Electron Microscopy Studies
   total water counta20 Å - OEC
 expt. typeresolution (Å)monomer Ibmonomer IIcsumIII
MD  1989  182d 
5B66 (43)SX1.85195719683925185179
5B5E (43)SX1.87181018143624185178
3WU2 (41)SX1.90150613782884184178
4UB6 (42)SX1.95137712132590176179
7D1T (67)Cryo-EM1.95122212102432174174
4IL6 (66) (Sr)eSX2.1010799582037169166
6W1O (47)XFEL (RT)2.0810819562037156156
6DHE (46)fXFEL (RT)2.0510829392021165155
6DHP (46)gXFEL (RT)2.0410729462018162159
6DHF (46)XFEL (RT)2.0810819362017165159
7RF1 (33)XFEL (RT)1.8910169441960165165
5GTI (44)XFEL2.509459661911162163
5GTH (44)XFEL2.508819001781158162
5WS5 (44)XFEL2.358098171626165166
7N8O (68)hCryo-EM1.936196171236156156
5TIS (45)XFEL (RT)2.255716081179116124
a

Average number of water molecules, data recorded every 50 ps. Water around the highly flexible regions (disordered in many crystallographic models) is excluded from the water count. The maximum and minimum total numbers of water molecules from the MD simulations are 2087 and 1897, respectively. If the flexible regions are included, then the average, maximum, and minimum numbers of water molecules around the complete PSII model from the MD simulations are 2050, 2153, and 1948, respectively. Water molecules with fractional occupancy in the experimental structures are counted as a single unit.

b

Chains with an upper letter case.

c

Chains with a lower letter case.

d

Maximum and minimum numbers of waters are 202 and 160, respectively.

e

Strontium-substituted OEC.

f

S1-enriched sample.

g

S0-enriched sample.

h

Mesophilic cyanobacterium Synechocystis sp. PCC 6803.

The number of resolved water molecules varies greatly among different crystallographic models, and non-negligible differences in water content may also exist between PSII monomers within the same crystal structure. The model of Tanaka et al. (43) (PDB ID: 5B66) exhibits the highest water content among all crystallographic models and is remarkably close in absolute terms to the water content predicted by the MD simulations. Importantly, this also implies that the MD simulations did not artificially result in overhydration of the computational model. Other crystallographic models derived from conventional synchrotron setups have lower water count. Structural models derived from XFEL-based crystallographic studies are the least hydrated. The model reported by Young et al. (45) (PDB ID: 5TIS) has the lowest water count, with apparent water content merely one third of that in the Tanaka et al. (43) model or in the present MD model, even though it does not have nominally the lowest resolution among XFEL models. The two cryo-EM structures are considerably different from each other in terms of water count. 7D1T (67) appears more hydrated than any XFEL model but still does not compare favorably with synchrotron-based crystallographic models.
The total hydration content is a global indicator, but information regarding internal water is more discerning as it relates to conserved channel systems. To probe this more specifically, we counted water molecules within a sphere of 20 Å radius centered at the OEC. The average number of water molecules within the OEC-centered spherical region from the MD simulations is 182 (the range spans a maximum of 202 and a minimum of 160 water molecules, which demonstrates the extent to which protein conformation and flexibility modulate hydration). Most of the conventional crystallographic models are in very good agreement with the MD simulations. Interestingly, the crystallographic models by Tanaka et al. (43) (5B66) and by Umena et al. (41) (3WU2), which showed large differences in the total hydration content (451 and 590 waters in the case of monomers I and II, respectively), yield nearly the same number of waters within the OEC-centered sphere. This shows that most unresolved water molecules in the 3WU2 model are close to the protein periphery and not in the interior.
The most interesting observation concerns again the XFEL-based crystallographic models. These present considerably lower water content even within the internal OEC-centered sphere, with the 5TIS model showing an unreasonably small number of water molecules. The low manifest water content at the donor side of PSII in XFEL-SFX models may be due to dehydration of the samples or to resolution limitations. However, the fact that the recent 7RF1 model by Hussein et al. (33) is reported to have an effective resolution rivaling the best synchrotron crystallographic models, yet suffers from a similarly low water count around the OEC as other XFEL-derived models, suggests that the diminished water count reflects a real departure of the samples from the physiological hydration state.
These results show that currently available XFEL-SFX models of PSII present an inadequate picture of the hydration state of the system, as a consequence of the severe dehydration of the samples. Crucially, these problems extend deep into the protein and even within the proximity of the OEC. Therefore, these models offer a precarious structural basis for inferences regarding the atomistic nature of water channels and may not be sufficiently reliable for extracting mechanistic information based on observed movements of water around the OEC site.

2.3. Architecture of Water Channels Around the OEC

In addition to the high oxidation potential generated at the reaction center, efficient catalytic action of the oxygen evolving complex requires channels for substrate water delivery, proton release, and efficient diffusion of dioxygen. In line with previous studies, our results show three major water channel systems in the lumenal side of PSII connecting the bulk phase with the OEC (see Figure 3). We follow recent convention and name them after the closest terminating atom of the OEC cluster, i.e., O1, O4, and Cl1 channel systems (the term “channel system” implies that they all consist of more than one water chain or branch). These approximately correspond to the “narrow”, “large”, and “broad” channels discussed in the past literature; for channel nomenclatures used in various studies, we refer the reader to the excellent summary by Hussein et al. (33) These systems remain hydrated throughout the simulation and no significant change in hydration content is observed within 20 Å around the OEC (Figure 2d). However, differences exist in terms of their length, width, tributaries (contributing minor channels that terminate at the protein–bulk interface), and presence of protein conformation-dependent transient channels. (14,16,20,25,27,30,69) We label the minor branches that contribute to the three major channel systems as E1–E9 (see Figure 3). In the following, we describe the key characteristics of each system.

Figure 3

Figure 3. Identification of water channels leading from the bulk toward the OEC, along with the proposal for the proton exit pathway associated with YZ. O1 (blue), O4 (red), and Cl1 (green) channel systems are colored uniquely for presentation purposes. Asterisks indicate positions of DMSO molecules found in the 5B66 crystallographic model. Dotted lines imply the transient nature of respective water channels due to protein conformational dynamics.

2.3.1. O1 Water Channel System

Channel system O1 contains in total four minor contributing channels (E1–E4, Figures 35), of which E1 and E2 remain hydrated and active in water exchange with the bulk throughout the simulation, whereas hydration of the other two channels E3 and E4 is dynamic and conformation-dependent. Specifically, which one is active depends on the conformational dynamics of PsbV-Tyr137, a C-terminal residue of the extrinsic PsbV protein. All four contributing channels merge at a region near D1-Glu329. The O1 channel system incorporates an internal water network associated with the redox-active D1-Tyr161 (YZ) residue that appears to connect with E4 but does not exchange waters with the bulk during our simulations.

Figure 4

Figure 4. Depiction of the E1 branch of the O1 channel system.

Figure 5

Figure 5. Depiction of the E2 branch of the O1 channel system. The superscript CTR denotes C-terminal residues.

The E1 channel is located at the interface between CP43 and PsbV. We observe that water can enter or exit E1 through two small water gates, labeled E1-1 and E1-2. In the case of E1-1, the incoming water makes first contact with the CP43-Glu300, CP43-Glu394, CP43-Arg391, and PsbV-Pro102 residues, whereas in the case of E1-2, contact is made in a region surrounded by PsbV-Arg31, PsbV-Gln34, PsbV-Tyr35, and CP43-Asn418 (Figure 4). Water molecules through both E1-1 and E1-2 merge at a junction consisting of a CP43-Glu83/PsbV-Lys103 salt-bridge and then pass through CP43-His398, CP43-Ala399, PsbV-Ser39, PsbV-Lys47 and CP43-Glu413 salt bridge, D1-Leu341, and CP43-Val410 to merge with the other channels (E2, E3, and E4) near D1-Glu329. The E1 is wider than the other contributors to the O1 channel system, and therefore, it contains more water molecules in the cross section. In this regard, it is also important to note that DMSO (dimethyl sulfoxide, used as cryoprotectant) is found inside this channel in the 5B66 crystallographic model (43) (Figure S5) and lies sufficiently close to the OEC, with its further penetration blocked by D1-Leu341, CP43-Val410, CP43-Leu401, and D1-Leu343. It is also noted that a glycerol molecule in the model by Umena et al. (41) is found to enter PSII through exactly the same cavity.
Water molecules in the E2 branch exchange through the interface created by residues D2-Glu343, CP47-Glu387, PsbU-Lys104 (C-terminal), D2-Leu352 (C-terminal), PsbV-Tyr137 (C-terminal), and D2-Arg348 (Figure 5). The C-terminal carboxylate groups of PsbU-Lys104, D2-Arg348, and D2-Leu352 form a triple salt-bridge-like interaction. The water in this channel also merges with other channels near D1-Glu329. It should be pointed out that the spatial region corresponding to this particular entry point overlaps with one of the entry points (E8) of the O4 water channel to be discussed in the following, i.e., the two channel systems are not entirely isolated from each other close to the interface.
The transient nature of the other two entry points, E3 and E4, is dependent on the conformational dynamics of PsbV-Tyr137 (C-terminal). In the case of E3, the incoming water first makes contact with the PsbF-Arg45 (C-terminal), PsbV-Lys30, and PsbJ-Leu40 (C-terminal) (Figure 6). The C-terminal carboxylate of PsbJ-Leu40 creates a triple salt-bridge-like interaction with the PsbV-Lys30 and PsbF-Arg45. Upon entry to this channel, the water passes through PsbV-Glu122, CP43-Ser416, PsbV-Trp130, PsbV-Lys129, and PsbV-Tyr137 and later merges with other channels at D1-Glu329. In the case of the E4 channel, we observed the formation of a linear water wire that is highly ordered so it might also serve in proton translocation (Figure 6). The incoming water in this case makes first contact with the PsbV-Tyr137 (C-terminal), PsbV-Lys129, and then with D1-Asp319 and D1-His304. Thereafter, the water passes through CP43-Ile414, CP43-Thr412, CP43-Glu413, and PsbV-Lys134 and merges with the other channels at D1-Glu329.

Figure 6

Figure 6. Depiction of the E3 and E4 branches of the O1 channel system, along with the YZ water network that converges to E4 close to the D1-Asp319 residue.

PsbV-Tyr137 populates two conformational states during the MD simulations, only one of which allows each one of the channels to become active (Figure 7), therefore functioning as a switch. This point regarding the conformational dependence of the activation of E3 and E4 is also reflected in the most hydrated crystal structure by Tanaka et al., (43) where only the E3 water channel is present, whereas E4 is absent (dehydrated, Figure S6). The hydration and formation of active water transport through E4 may have mechanistic consequences for the function of the OEC because the end-point of the E4 channel toward the bulk interacts with D1-Asp319, which is connected to the YZ(D1-Tyr161)-His190-Asn298 triad through an extensive hydrogen bond network of waters and residues.

Figure 7

Figure 7. Role of the PsbV-Tyr137 residue in gating the E3 and E4 channels by adopting two distinct orientations.

It is important to note that the YZ water system does not exchange waters with the bulk. The water molecules inside this channel are retained throughout the MD simulations, as in previous reports (25) and thus may only serve to establish a robust hydrogen bonding network for proton translocation upon YZ oxidation, presumably during one of the S-state transitions. (70,71) A peculiar part of the YZ water network is a pocket that is almost exclusively lined by asparagine residues (indicated as “Asn pocket” in Figure 3). (70) It has been speculated that the associated hydrogen bond network might serve for proton storage, stabilizing the proton via extensive hydrogen bonding, (25,72) potentially assisted by Asn tautomerization. (70) If a proton could also exit via the YZ branch, (70) then this may occur only when the E4 terminal is active, since this enables entry and exit of water necessary for proton discharge into the bulk. In this scenario, by controlling the state of E4, PsbV-Tyr137 may play a role in proton transfer from YZ. A recent site-directed mutagenesis study by Xiao et al. (73) showed the importance the PsbV-Tyr137 in regulating oxygen evolution and stabilizing other extrinsic proteins. Based on their results, it was proposed that PsbV-Tyr137 is crucial for the stable binding of the PsbV protein and for facilitating proton egress from YZ. Apart from PsbV-Tyr137, we also observe that PsbV-Lys134 makes an important salt-bridge interaction with the D1-Glu329 and D2-Leu352 (C-terminal). This particular region is important as all small tributaries of the O1 channel system merge here. Therefore, we conclude that the extrinsic PsbV protein is critical for the proper functioning and regulation of the entire O1 water channel system.
Recent site-directed mutagenesis experiments on the terminal residues of the E4 channels further supports the presents finding. Zhu et al. observed significantly decreased oxygen evolving activity (63–91% compared to the wild-type) in samples with mutations of D1-Arg323, D1-Asn322, D1-Asp319, and D1-His304. (74) These residues lie at the terminus of the E4 channel. In addition, it was found that the PsbV protein was lost in these mutants. (74) Overall, these experimental observations combined with the present computational results further strengthen the argument regarding the role of these key residues of the E4 channel in proton egress from the OEC, and they support the role of PsbV in regulating oxygen evolution.

2.3.2. Cl1 Water Channel System

This water channel system leading to the Mn4 atom of the OEC has three contributing branches (E5, E6, and E7), which remain functional throughout the MD simulations. E5 has a wider cross section and the shortest distance to the OEC and lies at the D1-D2-PsbO interface (Figure 8). The entry of water begins at the junction of the PsbO-Glu114, PsbO-His228, and PsbO-His231. Further progression leads to interaction with the D2-Glu310, PsbO-Arg115, D1-Tyr73, PsbO-Arg152, D1-Glu65, D1-Asp59, D2-Glu312 and D1-Arg334 salt-bridge, D1-Asp61, D2-Lys317, and D1-Asn181, and then finally toward the Mn4 site of the OEC. As expected from the large width of this channel, DMSO is found penetrating through this channel in the 5B66 crystal structure.

Figure 8

Figure 8. Depiction of the E5, E6, and E7 branches of the Cl1 channel system.

The E6 tributary is situated very close to the origin of E5. Water through this channel enters the protein scaffold near the interface between the carboxylate clusters PsbO-Asp222, PsbO-Asp223, PsbO-Asp224, and PsbO-Lys188. This water channel merges with E5 in the region near D1-Glu65 and PsbO-Arg152. The channel corresponding to E7 is a subsidiary of E6 and originates at the interface between the PsbO, PsbU, and CP47. In our simulations, we observed that water inside this branch can enter from multiple directions due to the very close proximity to the bulk. Water enters the protein matrix near the interface of PsbO-Arg189 and PsbU-Asn11 and then passes through PsbO-Ala185, PsbO-Arg162, and PsbO-Thr153. The E7 channel merges with E6 near the PsbO-Asp224 and PsbO-Lys188 (Figure 8).
We also observe a transient branch in the Cl1 channel system, which becomes active for certain time intervals of our MD simulation. Close inspection reveals that opening and closing of this channel is determined by rotation of the D1-Val67 side chain (Figure 9). The open conformation leads to filling with water toward the hydrophilic region near D1-Asp61. However, given that this transient channel has no direct connectivity to the bulk, it is not clear at the moment what role this channel might play in the regulation and function of the OEC. Interestingly, the water wire in this channel terminates near the chlorophyll ChlD1 of the reaction center, i.e., the pigment considered to be the primary electron donor of PSII. (75−78)

Figure 9

Figure 9. Transient water wire formation that is dependent on the conformation of D1-Val67: open (left) and closed (right) configurations determined by the side-chain rotation of D1-Val67.

2.3.3. O4 Water Channel System

The O4 system is the longest of the OEC-associated channel systems (Figure 10). In total, we observe two tributaries connecting it to the bulk, E8 and E9. Both entry points remained hydrated throughout the MD simulations. E8 is at the interface between the PsbU and CP47 proteins. The water exchanging through E8 first interacts with CP47-Lys389, PsbU-Glu23, and PsbU-Gln37. Thereafter, water passes through the region around PsbU-Tyr21, PsbU-Thr30, and CP47-Glu387 and subsequently toward the salt bridge formed by PsbU-Asp96 and CP43-Lys339. Further residues defining the channel are D2-Asn350, CP43-Pro334, D1-Asn335, and CP43-Leu337 toward the O4 atom of the OEC. The water between the PsbU-Asp96 and CP43-Lys339 salt bridge and the O4 forms a linear water wire consisting of up to 10 water molecules. DMSO solvent can be seen at the entrance of this channel (near PsbU-Tyr21) in the crystallographic model of PSII by Tanaka et al. (43) The E9 branch is located at the PsbU–PsbO interface, beginning around a salt bridge formed by PsbU-Glu93 and PsbO-Lys123, PsbO-Asn124 and PsbU-Thr89. A DMSO molecule is at the periphery of this channel in the 5B66 model. Water molecules through both the E8 and E9 branches merge at a region near the PsbU-Asp96 and CP43-Lys339 salt bridge. It is noted that in contrast to crystallographic models, (41−43) the MD results do not show any “break” in the continuity of waters and hence in the connectivity between the OEC and the bulk through the O4 channel. Such breaks observed in crystallographic models may therefore be artifacts of dehydration and not genuine structural features of the enzyme in its physiological state.

Figure 10

Figure 10. Depiction of the O4 channel system. The E8 and E9 branches merge close to the PsbU-Asp96/CP43-Lys339 pair.

2.3.4. Overview and General Remarks on Donor-Side Channel Systems

It is clear from the results and discussion above that all major channels leading from the bulk toward the OEC have multiple entry points. The origin of all water channels described here lies in the interface between the core polypeptides (i.e., D1, D2, CP43, and CP47) and the extrinsic proteins (PsbU, PsbV, and PsbO). The entry points for the water in the Cl1 channel system lie at the PSII dimer interface, whereas those of the O1 and O4 channel systems are away from the dimer interface, and hence PSII dimerization should not affect them. As we observe that a few of the channels originate around the conformationally flexible C-terminal residues (especially for the O1 channel system) and also involve salt-bridge interactions between core polypeptides and extrinsic proteins, the flexibility of such residues and ion-pairs can determine the opening or closing of channels or even the amount of water flux. Therefore, it is important to derive information on the channels from a dynamic treatment rather than from simple inspection of static crystallographic models, which typically have unresolved and disordered regions close to terminal residues.
Among the channels described above, we reported specific regions that contain highly ordered waters. The water molecules in these water “wires” do exchange with the bulk during our simulations, with the exception (at least within the time scale employed in this work) of the waters associated with YZ (Figure 6). The gated E4 channel, which has not been previously described in atomic detail, gains particular significance in this respect because it provides a structural basis for the possibility of proton translocation from the YZ site. Ordered waters reflect a highly organized hydrogen bonding network that is typically associated with Grotthuss-like proton transfer. This is the case for the ordered water molecules of the O4 network that are close to the OEC and that have been discussed as a possible pathway for proton release to the bulk. (23,79,80) This is most plausible if O4 is protonated in the S0 state of the OEC, (79) in which case, the proton would be released through the O4 network during the S0 → S1 transition. However, other studies favor O5 as the unique protonated bridge in the S0 state. (81−83) The question of proton release is even more significant for the later stages of the catalytic cycle, the S2 → S3 transition that prepares the cluster for oxygen evolution and the S3 → S0 transition that evolves O2 and resets the catalyst to its lowest oxidation level. Here, we note a recent study by Noguchi and coworkers (84) who investigated the effect of the D1-Glu65Ala mutation and suggested that the Cl1 channel serves as a single proton exit pathway in the S3 → S0 transition, whereas proton transfer in the S2 → S3 transition occurs through multiple pathways. Attribution of proton egress functionality to the Cl1 channel is in line with several other experimental and theoretical studies. (22,30,33,85−89) On the other hand, the conclusion regarding the nonspecificity of proton release in the S2 → S3 transition (84) mirrors the suggestion by Gunner and coworkers that all pathways are practically interconnected in the vicinity of the OEC, such that a proton released from any site of the cluster can in principle access all paths. (90) The complexity of these events and the persisting uncertainties regarding the precise structures of intermediates, particularly in the S3 state of the OEC, (47,70,91−95) pose continuing challenges for the detailed molecular understanding of proton transfer and associated channel specificity.
Conversely to the ordered water networks, high water mobility has been assumed to be beneficial for substrate delivery and in this case, the O1 channel system (which does indeed exhibit the highest rate of water exchange also in our simulations) has been associated with the delivery of water to the OEC for this reason. (33) Our simulations confirm relative differences in water mobility among these channels. However, this information does not allow us to assign functional roles because substrate binding and OEC catalytic cycling are strictly not incorporated in our simulations. Nevertheless, we would like to note that correlations of water mobility with function (i.e., the hypothesis that low mobility is more consistent with proton transfer whereas high mobility is more consistent with substrate delivery) are not self-evident. For example, water delivery may need to be precise more than it needs to be fast. This would be in line with the paramount importance of controlling reactivity at the highly oxidizing donor side of the PSII complex (9) whose overall chemistry is kinetically limited by the acceptor-side reactions (reduction of the terminal electron-acceptor plastoquinone QB). In this sense, the O4 channel may not yet be excluded as a water-delivery system; in fact, it is favored for this role by multiple steered molecular dynamics simulations of water permeation energetics (20) and by several studies that identify the O4 and Mn4 sites of the OEC as most accessible to small polar molecules such as ammonia and methanol. (34,35,96−104) In conclusion, assignments of functional roles for the donor-side channels, particularly in relation to S-state progression, remain uncertain and will require further studies to be reliably established.

2.4. Architecture of Water Channels on the Acceptor Side of PSII

More extensive differences in hydration with respect to available crystallographic models are identified at the stromal (acceptor) side of PSII. This contains the redox-active plastoquinones, the nonmobile primary electron acceptor QA, and the mobile terminal acceptor QB responsible for transporting reducing equivalents further along the photosynthetic chain to cytochrome b6f. A non-heme iron interfaces the QA and QB sites. The role of the iron-bound (bi)carbonate remains a subject of active research, but it is expected that bicarbonate and water are implicated in the protonation of the terminal acceptor QB. (105,106) Unlike the OEC site that is deeply buried within the protein matrix, the non-heme iron site is situated at the solvent-accessible stromal region; therefore, the hydration level in crystallographic models of this region is always and consistently lower compared to the lumenal region. It is emphasized that internal water molecules in the stromal side of the protein complex are far less likely to be conserved compared to internal water in the lumenal side that is capped by a host of extrinsic proteins. (14)
Our MD simulations lead to three crucial observations: (a) the stromal side of PSII is more hydrated than implied by even the highest-quality crystallographic models; (b) there are three well-organized water channel systems (labeled A, B, and C in the following, see Figure 11) connecting the bulk with the non-heme iron site, with multiple water entry/exit points; and (c) specific channels leading from the bulk toward the QB pocket are gated by protein conformational changes (side-chain motion and loop dynamics). Just like the channels observed around the OEC, the channels in the acceptor side differ from each other in their respective width, water influx, and local protein conformational dependence.

Figure 11

Figure 11. Schematic figure showing the three water channel systems connecting the non-heme iron site of PSII with the stromal bulk. Asterisks indicate positions of DMSO molecules found in the 5B66 crystallographic model. Dotted lines imply the transient nature of respective water channels due to protein conformational dynamics.

2.4.1. Channel A

This is the widest and shortest channel on the acceptor side. It remains hydrated throughout the simulation, and water can exchange easily between protein and bulk. The channel has two entry points for water, labeled A1 and A2 (Figures 11 and 12). In the case of A1, the water first interacts via hydrogen bonding with the backbone carbonyl of D2-Pro237, D2-Thr238, D2-Gln239, and D1-Tyr-246. Upon entry, the water establishes hydrogen bonding interaction with D1-Ser268. Aided by the large flexibility of its side-chain, D1-Ser268 “guides” the water between the bicarbonate group (HCO3) and D1-Tyr-246. This particular region is special because at a given time, a single water can simultaneously hydrogen-bond with the D1-Ser268, bicarbonate, D1-Tyr-246, and D1-Glu244. Water molecules in the region have large residence times, which may have important implications for the redox and proton transport events at this site. Interestingly, the existence of this channel might be inferred by the 5B66 crystallographic model of Tanaka et al. (43) (a DMSO molecule is found at the entrance of this channel, see Figure S7); however, it is dehydrated compared to the MD simulations. In the “worst-case” of the XFEL-SFX models, no water molecules are present and hence the existence of the channel is not obvious.

Figure 12

Figure 12. (a) Identification of the A1 water network leading from the bulk to the non-heme iron site; (b) A2 channel in the closed state, channel A1 is also depicted; (c) A2 channel in the open state, merging with channel A1.

The A2 branch is transient in nature and depends on the local conformational dynamics of the protein. Water entering A2 interacts with the D2-Asp225 and CP43-Arg461 salt bridge, D2-Glu241, D2-Gln239, D2-Thr243, and D1-Arg269 (Figure 12). Based on our simulations, the region around D2-Gln239 acts as a gate for this channel. Owing to its high side chain mobility, it adopts open and closed conformations that allow water to merge toward the A1 branch near D1-Ser268 (Figure 12 and S8). D2-Phe235 is another important residue controlling this specific channel, since its swaying motion allows modulation of the width of the channel, regulating water passage. In the 5B66 crystal structure, this channel is dehydrated due to the closed conformation of side-chain of D2-Gln239 (Figure S7). A few waters can be seen at the entrance of this channel in the crystallographic model, resembling the hydration structure predicted by our MD simulations when the channel is closed. We will not speculate on the possible functional relevance of this channel, but we note that a site-directed mutagenesis study (107) involving the D1-Arg269Gly substitution demonstrated impaired oxygen evolution and photosynthetic growth.

2.4.2. Channel B

This channel provides a gateway for bulk water to access the NHI site through an extensive hydrogen bond network. This water channel has two obvious entry points, labeled B1 and B2 (Figure 13). In case of B1, the water interacts with D1-Glu242 and CP43-Glu464, while a series of hydrogen bonding interactions are facilitated by D2-Ser245, the peptide -NH groups of D2-Val247 and D2-Met246, backbone carbonyls of D1-Gln241, D1-Tyr237, and the D2-Glu242/D2-Lys264/D1-Glu244 triple salt bridge (Figure 13). We observe that exchange of water from this channel to the NHI site is highly dependent on the conformational flexibility of the D2-Glu242 and D2-Lys264 salt bridge. For the majority of the simulation time, water exchange does not occur. However, the connectivity of the channel to the NHI site through hydrogen bonding remains intact. This channel can also be seen in several high-resolution crystallographic models of PSII. (33,41,43)

Figure 13

Figure 13. B1 (left) and B2 (right) water branches connecting the bulk with the NHI site.

Branch B2 is slightly shorter in length compared to B1. Water entering through B2 first interacts with D1-Glu243 and D1-Glu236 and thereafter establishes hydrogen bonding with the side-chains of D1-Gln241 and D1-Thr245. The side-chain of the D1-Gln241 is further hydrogen bonded to the water molecules of the B1 branch. B2 remains hydrated throughout our MD simulations. This particular channel can also be seen in crystal structures by Tanaka et al. (43) and Umena et al. (41) (Figure S9). A DMSO molecule can also be seen at the entrance of this channel in the 5B66 model.

2.4.3. Channel C and the QB Pocket

This channel is unique and has not been previously resolved crystallographically. Water present in this channel interacts with the NHI site through the QB hydrophobic pocket, and hence, it is expected to play a crucial role in QB protonation. Water inside this channel can exchange with the bulk from two sides: (a) through channel A and (b) through an area between D1-Ile248, D1-Asn266, D2-Pro237, and D2-Phe235 (Figure 14).

Figure 14

Figure 14. (a) Entry of water into channel C blocked by the hydrophobic side chains of D1-Ile248 and D1-Leu271; (b) formation of water wire connecting the bicarbonate site through the QB pocket with the bulk; (c) QB pocket water trapped because exit is restricted by the side chain conformation of D1-Asn266.

The entry of water through both endpoints is gated by the local conformation dynamics of the protein. Specifically, the entry of water to channel C through channel A depends on the side chain rotations of D1-Leu271 and D1-Ile248: the cavity is open when the side-chains are facing away from each other; otherwise, it is closed (Figure 14). We observe that the gating by these residues in the QB pocket is the strongest determinant for the functionality of this channel. The QB pocket can only hold a maximum of two water molecules at a time.
With the knowledge about the existence of this channel derived from our MD simulations, we revisited the crystallographic models and identified a structural “trace” of this previously unknown channel in the crystallographic model of Umena et al. (41) (but not in the model of Tanaka et al. (43)) where D1-Ile248 is disordered and present in two conformations, which correspond to the open and closed conformations of our computational model with occupancies of 70 and 30%, respectively (Figure S10).
A crucial point concerns the hydration of the QB pocket. We observed that the presence of two water molecules inside the QB cavity was initially predicted by our 3D-RISM calculations. Subsequent MC/MD calculations never removed these waters from the cavity. To further confirm that the hydration of this cavity is not an artifact of the initial system preparation, we performed MD simulations after removing these waters. We observed that the cavity is filled up again with water within the first 5 ns of the MD simulation, confirming that the presence of these two water molecules represents a physiological hydration state of the system.
The entry and exit of the water through D1-Ile248, D1-Asn266, D2-Pro237, and D2-Phe235 regions depend on the motion of two loops, the first one consisting of D1-Asn267, D1-Asn266, D1-Phe265, and D1-Ser264 and the second one consisting of D2-Gln239, D2-Thr238, D2-Pro237, D2-Asn236, and D2-Phe235. These two loops were found to be highly flexible in our MD simulations (Figure S11). In addition, the side chain of D1-Asn266 was also found to play a crucial role in the opening and closing of this channel from this particular end-point. This implies that even if water enters through channels A and B into the QB pocket, it may not leave until the other end is open. This also leads to large residence times of the waters in the QB pocket. Interestingly, the exit of the waters through these loops is at the very origin of the A1 branch. It is important to highlight the special role of the loop consisting of D2-Gln239, D2-Thr238, D2-Pro237, D2-Asn236, and D2-Phe235, since it forms the gateway for A1 and subsequently controls movement of water through C and A2. Although this particular loop region was found to be in the open state in many crystal structures, no water molecules were resolved. D1-His252 is present at the entrance of this channel; this residue is suggested to be critical (108) in providing the first proton to the QB through D1-Ser264. (109)

2.4.4. Hydrogen Bonding Network Around the NHI Site

The hydrogen bonding network between the protein residues and channel waters is dynamic yet retains a high and clearly identifiable structure and organization. As this region is close to the protein–bulk interface, it is subject to high perturbation from the bulk. It is also evident that the side chains of many residues around the NHI are highly flexible compared to more deeply buried residues. Perhaps counterintuitively, side-chain flexibility is found to be crucial for maintaining organization in water channels leading to the NHI site because by easy rotation of, for example, D1-Ser268, the residue can always either establish a hydrogen bonding interaction with the incoming water or stabilize (and hence maintain the structure of) the hydrogen bonded water tetramer near the bicarbonate. We note that recently targeted mutagenesis by Forsman and Eaton-Rye (110) highlighted the importance of D1-Ser268 in conjunction with the bicarbonate for proton transfer events in the formation of QBH2. The importance of D1-Ser268 is also highlighted by the site-directed mutagenesis studies of Alfonso et al., (111) who observed that serine to proline substitution leads to a decrease in the electron transfer rate between QA and QB and stabilization of S2QB and S3QB states.
The triple salt bridge consisting of D1-Glu244, D2-Lys264, and D2-Glu242 is significant in terms of directly interacting with the incoming waters from channels A and B and stabilizing them through extensive hydrogen bonding. This structural observation derived from the MD simulations may relate to experimental results of site-directed mutagenesis (112) on D1-Glu244 and D2-Tyr246, which resulted in impaired oxygen evolution, implying that these residues maintain an important hydrogen bonding network critical for the smooth function of the acceptor side. Similarly, recent site-directed mutagenesis study by Khaing et al. (113) on D2-Tyr244 showed impaired electron transfer between QA and QB, including disruption in the PSII assembly and in the back-reaction with the S2 state of the OEC. In addition, Fourier transform infrared spectroscopy studies by Hienerwadel and Berthomieu (114) documented a hydrogen bond network from the non-heme iron toward the QB pocket. Based on our MD results, we do see such a strong network, which involves the non-heme iron, D1-Glu244, D1-Ser268, D1-Tyr246, and two water molecules in the QB pocket. Similarly, Takahashi et al. (115) claimed that one of the two tyrosines participates in a direct hydrogen bonding interaction with the bicarbonate ligand. Based on our simulations, D2-Tyr244 is serving as hydrogen bond donor to both the bicarbonate and the nearby D2-Met246.

2.4.5. Role of Hydration in QB Protonation in PSII and the Bacterial Reaction Center

PSII and the bacterial reaction center (bRC) share striking similarity in the architecture around the quinones and non-heme iron site but with some key important structural differences (Figure 15). (116) This is clearly reflected in the electron transfer kinetics, for example, the rates of first and second electron transfers from QA to QB are different in bRC and PSII. The first electron is transferred within 100 μs in bRC, (117) whereas it takes 400 μs in the case of the PSII. (118) The second electron transfer event is much slower, at 1 ms in bRC (117) and 0.6–0.8 ms in PSII. (118) In the reaction center of purple bacteria, after the first electron transfer from the primary acceptor ubiquinone QA to the terminal acceptor ubiquinone QB, the first proton is known to be delivered through L-Ser223, which in return receives a proton from L-Asp213 (Figure 15). After re-reduction of QB by QA, the second proton may be provided by the L-Glu212 residue. (119,120) Sugo et al. (121) suggest that the second proton is derived from L-His190 instead, whereas L-Glu212 is involved in water-mediated proton transfer to the deprotonated L-His190. Assuming that PSII operates in a similar fashion, it is speculated that the first proton delivered to QB is derived from D1-Ser264, which in return receives the proton from D1-His252 that has access to the bulk. Some experimental and theoretical studies suggest instead that the second proton is derived from D1-His215. (105,106,116)

Figure 15

Figure 15. Differences in the protein environment around the analogous quinones in (a) the Photosystem II (PDB ID 3WU2) and (b) the bacterial reaction center (PDB ID 3I4D).

The large time-scales of the sequential electron and proton transfer processes suggest strong coupling between the conformational changes of the protein and redox events. Water can be functionally important given the proximity of the non-heme iron site to the bulk in both the PSII and the bRC. Interestingly, the role of overall protein hydration in the QA to QB electron transfer kinetics has been discussed in the case of the bRC. (56,57) Again, it highlights the significance of water in providing functional flexibility to the protein scaffold and established its mechanistic role in redox events around the QB site. Similarly, the role of hydration in terms of the electron transfer kinetics in the acceptor side has been demonstrated also for PSII. (60,61) Dehydration leads to impaired electron transfer from the QA to QB or no electron transfer at all. Therefore, correctly and fully hydrated PSII models are the key for understanding the system’s overall structure and function.
The QB pocket waters discussed above in connection to channel C have not been previously identified in existing crystal structures of PSII. Given their proximity to the bicarbonate and QB, and in view of the connectivity to the bulk, they may be involved in proton transfer within the QB cavity. Based on the MD simulations and previous results from various experimental and computational studies, the QB pocket waters can play two roles: (a) mediate proton transfer from the bicarbonate or bulk to QB, and (b) mediate proton transfer to the deprotonated D1-His215. In addition to the role of the water channels in proton transfer, a recent work by Fantuzzi et al. (122) demonstrated the role of the water channels in dioxygen diffusion toward the non-heme site. Assuming that D1-His215 transfers the second proton to QB and becomes doubly deprotonated, the QB pocket water molecules can transfer a proton to D1-His215. In this case, the proton can enter from the bulk or from the bicarbonate itself. Ishikita and co-workers (109) postulated the presence of the QB pocket water for the protonation of the D1-His215 anion. (123) In addition, the second proton transfer can also be initiated by the QB pocket waters due to their close proximity; however, this would require some degree of rotation of the QB head-group. This particular proposal remains to be investigated, and it would be interesting to see how the electron transfer events are coupled to the conformational changes in the protein around the QB region.

3. Conclusions

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In this work, we presented large scale molecular dynamics simulations of membrane-bound Photosystem II to understand its hydration structure, employing a coherent protocol for the generation of a fully hydrated system before production level simulations. Three aspects were targeted: (a) overall hydration of the PSII and comparison with the recent and previously known crystallographic models, (b) water channel architecture around the OEC, and (c) water channel architecture at the acceptor side of PSII.
The hydration count predicted by our MD simulations best matches with the structure reported by Tanaka et al., (43) which represents the highest-resolution synchrotron-based crystallographic model of PSII. Other crystallographic models display lower water counts. At the extreme, recent models derived from XFEL crystallography were found to be severely dehydrated, not only in the periphery of PSII but also internally and even around the OEC.
Our results map in great detail the three water channel systems connecting the OEC with the bulk. We document the multiple water entry/exit points of these channels and identify a number of previously unknown transiently formed water pathways. Protein conformational dynamics plays a key role in the functionality of several channels. Especially, we highlight the role of PsbV-Tyr137 in conformationally gating the water channels and its possible involvement in proton egress from the YZ network. Almost all water channels are interfaced between the core proteins and extrinsic proteins, underlining the role of extrinsic protein in regulating water access to the OEC and proton egress to the lumen.
On the acceptor side of PSII, we identify an extensive and well-organized water channel system, which typically remains dehydrated in most experimental structures. An important discovery is a previously unknown channel that is responsible for hydration of the QB pocket and hence may play an important role in proton translocation around the non-heme iron site or even directly in the protonation of the terminal acceptor QB.
The present study offers new insight into the equilibrium water content of PSII and the dynamic nature of its static and transient water channels. The fully hydrated structures of PSII, provided as an open-data collection of supplementary PDB files, can be used for evaluating the hydration content of crystallographic models and serve as starting point for future investigations into the precise mechanistic role of water in donor- and acceptor-side processes.

Supporting Information

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The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/jacs.2c09121.

  • Detailed description of methodology including system setup, parameterization, and protocols used for the molecular dynamics simulations; analysis of molecular dynamics trajectories (Figures S1–S3); distribution of 3D-RISM positioned water in the QB cavity (Figure S4); and comparisons with crystallographic models and depiction of conformational flexibility of key residues (Figures S5–S11) (PDF)

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Author Information

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  • Corresponding Author
  • Author
    • Abhishek Sirohiwal - Max-Planck-Institut für Kohlenforschung, Kaiser-Wilhelm-Platz 1, 45470 Mülheim an der Ruhr, GermanyPresent Address: Department of Biochemistry and Biophysics, Arrhenius Laboratory, Stockholm University, 10691 Stockholm, Sweden (A.S.)Orcidhttps://orcid.org/0000-0002-4073-7627
  • Funding

    Open access funded by Max Planck Society.

  • Notes
    The authors declare no competing financial interest.

    Original PDB files from the MD simulations selected to depict representative hydration states of the water channels identified in the present work are provided as an open-access data set hosted by the Open Research Data Repository of the Max Planck Society at https://doi.org/10.17617/3.B2AKNU.

Acknowledgments

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The authors acknowledge support by the Max Planck Society and thank the Max Planck Computing and Data Facility (MPCDF, Garching, Germany) for computational resources.

References

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This article references 123 other publications.

  1. 1
    Shevela, D.; Björn, L. O.; Govindjee. Photosynthesis: Solar Energy for Life; World Scientific: Singapore, 2017, p 204.
  2. 2
    Blankenship, R. E. Molecular Mechanisms of Photosynthesis; 2nd ed.; Wiley: Chichester, 2014, p 312.
  3. 3
    Photosystem II. The Light-Driven Water:Plastoquinone Oxidoreductase; Wydrzynski, T.; Satoh, K., Eds.; Springer: Dordrecht, 2005; Vol. 22, p 786.
  4. 4
    Barber, J. Photosystem II: the water splitting enzyme of photosynthesis and the origin of oxygen in our atmosphere. Q. Rev. Biophys. 2016, 49, e14  DOI: 10.1017/S0033583516000093
  5. 5
    Krewald, V.; Retegan, M.; Pantazis, D. A. Principles of Natural Photosynthesis. Top. Curr. Chem. 2016, 371, 2348,  DOI: 10.1007/128_2015_645
  6. 6
    Shen, J.-R. The Structure of Photosystem II and the Mechanism of Water Oxidation in Photosynthesis. Annu. Rev. Plant Biol. 2015, 66, 2348,  DOI: 10.1146/annurev-arplant-050312-120129
  7. 7
    Pantazis, D. A. In Hydrogen Production and Energy Transition; Van de Voorde, M., Ed.; De Gruyter: 2021; Vol. 1, p 427468.
  8. 8
    Junge, W. Oxygenic photosynthesis: history, status and perspective. Q. Rev. Biophys. 2019, 52, e1  DOI: 10.1017/S0033583518000112
  9. 9
    Pantazis, D. A. Missing Pieces in the Puzzle of Biological Water Oxidation. ACS Catal. 2018, 8, 94779507,  DOI: 10.1021/acscatal.8b01928
  10. 10
    Cox, N.; Pantazis, D. A.; Lubitz, W. Current Understanding of the Mechanism of Water Oxidation in Photosystem II and Its Relation to XFEL Data. Annu. Rev. Biochem. 2020, 89, 795820,  DOI: 10.1146/annurev-biochem-011520-104801
  11. 11
    Ho, F. M. Uncovering channels in photosystem II by computer modelling: current progress, future prospects, and lessons from analogous systems. Photosynth. Res. 2008, 98, 503522,  DOI: 10.1007/s11120-008-9358-2
  12. 12
    Ho, F. M.; Styring, S. Access Channels and Methanol Binding Site to the CaMn4 Cluster in Photosystem II Based on Solvent Accessibility Simulations, with Implications for Substrate Water Access. Biochim. Biophys. Acta, Bioenerg. 2008, 1777, 140153,  DOI: 10.1016/j.bbabio.2007.08.009
  13. 13
    Ho, F. M. In Molecular Solar Fuels; Wydrzynski, T. J.; Hillier, W., Eds.; The Royal Society of Chemistry: Cambridge, 2012, p 208248.
  14. 14
    Linke, K.; Ho, F. M. Water in Photosystem II: Structural, Functional and Mechanistic Considerations. Biochim. Biophys. Acta, Bioenerg. 2014, 1837, 1432,  DOI: 10.1016/j.bbabio.2013.08.003
  15. 15
    Nakamura, S.; Ota, K.; Shibuya, Y.; Noguchi, T. Role of a Water Network around the Mn4CaO5 Cluster in Photosynthetic Water Oxidation: A Fourier Transform Infrared Spectroscopy and Quantum Mechanics/Molecular Mechanics Calculation Study. Biochemistry 2016, 55, 597607,  DOI: 10.1021/acs.biochem.5b01120
  16. 16
    Bondar, A.-N.; Dau, H. Extended Protein/Water H-bond Networks in Photosynthetic Water Oxidation. Biochim. Biophys. Acta, Bioenerg. 2012, 1817, 11771190,  DOI: 10.1016/j.bbabio.2012.03.031
  17. 17
    Murray, J.; Barber, J. In Photosynthesis. Energy from the Sun; Allen, J.; Gantt, E.; Golbeck, J.; Osmond, B., Eds.; Springer Netherlands: 2008, p 467470,  DOI: 10.1007/978-1-4020-6709-9_105 .
  18. 18
    Gabdulkhakov, A.; Guskov, A.; Broser, M.; Kern, J.; Müh, F.; Saenger, W.; Zouni, A. Probing the Accessibility of the Mn4Ca Cluster in Photosystem II: Channels Calculation, Noble Gas Derivatization, and Cocrystallization with DMSO. Structure 2009, 17, 12231234,  DOI: 10.1016/j.str.2009.07.010
  19. 19
    Vassiliev, S.; Comte, P.; Mahboob, A.; Bruce, D. Tracking the flow of water through photosystem II using molecular dynamics and streamline tracing. Biochemistry 2010, 49, 18731881,  DOI: 10.1021/bi901900s
  20. 20
    Vassiliev, S.; Zaraiskaya, T.; Bruce, D. Exploring the Energetics of Water Permeation in Photosystem II by Multiple Steered Molecular Dynamics Simulations. Biochim. Biophys. Acta, Bioenerg. 2012, 1817, 16711678,  DOI: 10.1016/j.bbabio.2012.05.016
  21. 21
    Vassiliev, S.; Zaraiskaya, T.; Bruce, D. Molecular Dynamics Simulations Reveal Highly Permeable Oxygen Exit Channels Shared with Water Uptake Channels in Photosystem II. Biochim. Biophys. Acta, Bioenerg. 2013, 1827, 11481155,  DOI: 10.1016/j.bbabio.2013.06.008
  22. 22
    Ishikita, H.; Saenger, W.; Loll, B.; Biesiadka, J.; Knapp, E.-W. Energetics of a Possible Proton Exit Pathway for Water Oxidation in Photosystem II. Biochemistry 2006, 45, 20632071,  DOI: 10.1021/bi051615h
  23. 23
    Takaoka, T.; Sakashita, N.; Saito, K.; Ishikita, H. pKa of a Proton-Conducting Water Chain in Photosystem II. J. Phys. Chem. Lett. 2016, 7, 19251932,  DOI: 10.1021/acs.jpclett.6b00656
  24. 24
    Sakashita, N.; Watanabe, H. C.; Ikeda, T.; Ishikita, H. Structurally conserved channels in cyanobacterial and plant photosystem II. Photosynth. Res. 2017, 133, 7585,  DOI: 10.1007/s11120-017-0347-1
  25. 25
    Sakashita, N.; Watanabe, H. C.; Ikeda, T.; Saito, K.; Ishikita, H. Origins of Water Molecules in the Photosystem II Crystal Structure. Biochemistry 2017, 56, 30493057,  DOI: 10.1021/acs.biochem.7b00220
  26. 26
    Sakashita, N.; Ishikita, H.; Saito, K. Rigidly hydrogen-bonded water molecules facilitate proton transfer in photosystem II. Phys. Chem. Chem. Phys. 2020, 22, 1583115841,  DOI: 10.1039/D0CP00295J
  27. 27
    Ogata, K.; Hatakeyama, M.; Sakamoto, Y.; Nakamura, S. Investigation of a Pathway for Water Delivery in Photosystem II Protein by Molecular Dynamics Simulation. J. Phys. Chem. B 2019, 123, 64446452,  DOI: 10.1021/acs.jpcb.9b04838
  28. 28
    Ghosh, I.; Khan, S.; Banerjee, G.; Dziarski, A.; Vinyard, D. J.; Debus, R. J.; Brudvig, G. W. Insights into Proton-Transfer Pathways during Water Oxidation in Photosystem II. J. Phys. Chem. B 2019, 123, 81958202,  DOI: 10.1021/acs.jpcb.9b06244
  29. 29
    Kaur, D.; Cai, X.; Khaniya, U.; Zhang, Y.; Mao, J.; Mandal, M.; Gunner, M. R. Tracing the Pathways of Waters and Protons in Photosystem II and Cytochrome c Oxidase. Inorganics 2019, 7, 14,  DOI: 10.3390/inorganics7020014
  30. 30
    Guerra, F.; Siemers, M.; Mielack, C.; Bondar, A.-N. Dynamics of Long-Distance Hydrogen-Bond Networks in Photosystem II. J. Phys. Chem. B 2018, 122, 46254641,  DOI: 10.1021/acs.jpcb.8b00649
  31. 31
    Saito, K.; Rutherford, A. W.; Ishikita, H. Mechanism of Tyrosine D Oxidation in Photosystem II. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 76907695,  DOI: 10.1073/pnas.1300817110
  32. 32
    Sirohiwal, A.; Neese, F.; Pantazis, D. A. Microsolvation of the Redox-Active Tyrosine-D in Photosystem II: Correlation of Energetics with EPR Spectroscopy and Oxidation-Induced Proton Transfer. J. Am. Chem. Soc. 2019, 141, 32173231,  DOI: 10.1021/jacs.8b13123
  33. 33
    Hussein, R.; Ibrahim, M.; Bhowmick, A.; Simon, P. S.; Chatterjee, R.; Lassalle, L.; Doyle, M.; Bogacz, I.; Kim, I.-S.; Cheah, M. H.; Gul, S.; de Lichtenberg, C.; Chernev, P.; Pham, C. C.; Young, I. D.; Carbajo, S.; Fuller, F. D.; Alonso-Mori, R.; Batyuk, A.; Sutherlin, K. D.; Brewster, A. S.; Bolotovsky, R.; Mendez, D.; Holton, J. M.; Moriarty, N. W.; Adams, P. D.; Bergmann, U.; Sauter, N. K.; Dobbek, H.; Messinger, J.; Zouni, A.; Kern, J.; Yachandra, V. K.; Yano, J. Structural dynamics in the water and proton channels of photosystem II during the S2 to S3 transition. Nat. Commun. 2021, 12, 6531,  DOI: 10.1038/s41467-021-26781-z
  34. 34
    Retegan, M.; Pantazis, D. A. Interaction of methanol with the oxygen-evolving complex: atomistic models, channel identification, species dependence, and mechanistic implications. Chem. Sci. 2016, 7, 64636476,  DOI: 10.1039/C6SC02340A
  35. 35
    Retegan, M.; Pantazis, D. A. Differences in the Active Site of Water Oxidation among Photosynthetic Organisms. J. Am. Chem. Soc. 2017, 139, 1434014343,  DOI: 10.1021/jacs.7b06351
  36. 36
    Vogt, L.; Vinyard, D. J.; Khan, S.; Brudvig, G. W. Oxygen-evolving complex of Photosystem II: an analysis of second-shell residues and hydrogen-bonding networks. Curr. Opin. Chem. Biol. 2015, 25, 152158,  DOI: 10.1016/j.cbpa.2014.12.040
  37. 37
    Weisz, D. A.; Gross, M. L.; Pakrasi, H. B. Reactive oxygen species leave a damage trail that reveals water channels in Photosystem II. Sci. Adv. 2017, 3, eaao3013  DOI: 10.1126/sciadv.aao3013
  38. 38
    Loll, B.; Kern, J.; Saenger, W.; Zouni, A.; Biesiadka, J. Towards complete cofactor arrangement in the 3.0 Å resolution structure of photosystem II. Nature 2005, 438, 10401044,  DOI: 10.1038/nature04224
  39. 39
    Ferreira, K. N.; Iverson, T. M.; Maghlaoui, K.; Barber, J.; Iwata, S. Architecture of the Photosynthetic Oxygen-Evolving Center. Science 2004, 303, 18311838,  DOI: 10.1126/science.1093087
  40. 40
    Murray, J. W.; Barber, J. Structural Characteristics of Channels and Pathways in Photosystem II Including the Identification of an Oxygen Channel. J. Struct. Biol. 2007, 159, 228237,  DOI: 10.1016/j.jsb.2007.01.016
  41. 41
    Umena, Y.; Kawakami, K.; Shen, J.-R.; Kamiya, N. Crystal Structure of the Oxygen-Evolving Photosystem II at a Resolution of 1.9 Å. Nature 2011, 473, 5560,  DOI: 10.1038/nature09913
  42. 42
    Suga, M.; Akita, F.; Hirata, K.; Ueno, G.; Murakami, H.; Nakajima, Y.; Shimizu, T.; Yamashita, K.; Yamamoto, M.; Ago, H.; Shen, J.-R. Native Structure of Photosystem II at 1.95 Å Resolution Viewed by Femtosecond X-ray Pulses. Nature 2015, 517, 99103,  DOI: 10.1038/nature13991
  43. 43
    Tanaka, A.; Fukushima, Y.; Kamiya, N. Two different structures of the oxygen-evolving complex in the same polypeptide frameworks of photosystem II. J. Am. Chem. Soc. 2017, 139, 17181721,  DOI: 10.1021/jacs.6b09666
  44. 44
    Suga, M.; Akita, F.; Sugahara, M.; Kubo, M.; Nakajima, Y.; Nakane, T.; Yamashita, K.; Umena, Y.; Nakabayashi, M.; Yamane, T.; Nakano, T.; Suzuki, M.; Masuda, T.; Inoue, S.; Kimura, T.; Nomura, T.; Yonekura, S.; Yu, L.-J.; Sakamoto, T.; Motomura, T.; Chen, J.-H.; Kato, Y.; Noguchi, T.; Tono, K.; Joti, Y.; Kameshima, T.; Hatsui, T.; Nango, E.; Tanaka, R.; Naitow, H.; Matsuura, Y.; Yamashita, A.; Yamamoto, M.; Nureki, O.; Yabashi, M.; Ishikawa, T.; Iwata, S.; Shen, J.-R. Light-Induced Structural Changes and the Site of O=O bond Formation in PSII Caught by XFEL. Nature 2017, 543, 131135,  DOI: 10.1038/nature21400
  45. 45
    Young, I. D.; Ibrahim, M.; Chatterjee, R.; Gul, S.; Fuller, F. D.; Koroidov, S.; Brewster, A. S.; Tran, R.; Alonso-Mori, R.; Kroll, T.; Michels-Clark, T.; Laksmono, H.; Sierra, R. G.; Stan, C. A.; Hussein, R.; Zhang, M.; Douthit, L.; Kubin, M.; de Lichtenberg, C.; Vo Pham, L.; Nilsson, H.; Cheah, M. H.; Shevela, D.; Saracini, C.; Bean, M. A.; Seuffert, I.; Sokaras, D.; Weng, T.-C.; Pastor, E.; Weninger, C.; Fransson, T.; Lassalle, L.; Bräuer, P.; Aller, P.; Docker, P. T.; Andi, B.; Orville, A. M.; Glownia, J. M.; Nelson, S.; Sikorski, M.; Zhu, D.; Hunter, M. S.; Lane, T. J.; Aquila, A.; Koglin, J. E.; Robinson, J.; Liang, M.; Boutet, S.; Lyubimov, A. Y.; Uervirojnangkoorn, M.; Moriarty, N. W.; Liebschner, D.; Afonine, P. V.; Waterman, D. G.; Evans, G.; Wernet, P.; Dobbek, H.; Weis, W. I.; Brunger, A. T.; Zwart, P. H.; Adams, P. D.; Zouni, A.; Messinger, J.; Bergmann, U.; Sauter, N. K.; Kern, J.; Yachandra, V. K.; Yano, J. Structure of Photosystem II and Substrate Binding at Room Temperature. Nature 2016, 540, 453457,  DOI: 10.1038/nature20161
  46. 46
    Kern, J.; Chatterjee, R.; Young, I. D.; Fuller, F. D.; Lassalle, L.; Ibrahim, M.; Gul, S.; Fransson, T.; Brewster, A. S.; Alonso-Mori, R.; Hussein, R.; Zhang, M.; Douthit, L.; de Lichtenberg, C.; Cheah, M. H.; Shevela, D.; Wersig, J.; Seuffert, I.; Sokaras, D.; Pastor, E.; Weninger, C.; Kroll, T.; Sierra, R. G.; Aller, P.; Butryn, A.; Orville, A. M.; Liang, M.; Batyuk, A.; Koglin, J. E.; Carbajo, S.; Boutet, S.; Moriarty, N. W.; Holton, J. M.; Dobbek, H.; Adams, P. D.; Bergmann, U.; Sauter, N. K.; Zouni, A.; Messinger, J.; Yano, J.; Yachandra, V. K. Structures of the Intermediates of Kok’s Photosynthetic Water Oxidation Clock. Nature 2018, 563, 421425,  DOI: 10.1038/s41586-018-0681-2
  47. 47
    Ibrahim, M.; Fransson, T.; Chatterjee, R.; Cheah, M. H.; Hussein, R.; Lassalle, L.; Sutherlin, K. D.; Young, I. D.; Fuller, F. D.; Gul, S.; Kim, I.-S.; Simon, P. S.; de Lichtenberg, C.; Chernev, P.; Bogacz, I.; Pham, C. C.; Orville, A. M.; Saichek, N.; Northen, T.; Batyuk, A.; Carbajo, S.; Alonso-Mori, R.; Tono, K.; Owada, S.; Bhowmick, A.; Bolotovsky, R.; Mendez, D.; Moriarty, N. W.; Holton, J. M.; Dobbek, H.; Brewster, A. S.; Adams, P. D.; Sauter, N. K.; Bergmann, U.; Zouni, A.; Messinger, J.; Kern, J.; Yachandra, V. K.; Yano, J. Untangling the sequence of events during the S2 → S3 transition in photosystem II and implications for the water oxidation mechanism. Proc. Natl. Acad. Sci. U. S. A. 2020, 117, 1262412635,  DOI: 10.1073/pnas.2000529117
  48. 48
    Sanchez-Weatherby, J.; Moraes, I. In The Next Generation in Membrane Protein Structure Determination; Moraes, I., Ed.; Springer International Publishing: Cham, 2016, p 7389.  DOI: 10.1007/978-3-319-35072-1_6 .
  49. 49
    Hellmich, J.; Bommer, M.; Burkhardt, A.; Ibrahim, M.; Kern, J.; Meents, A.; Müh, F.; Dobbek, H.; Zouni, A. Native-like Photosystem II Superstructure at 2.44 Å Resolution through Detergent Extraction from the Protein Crystal. Structure 2014, 22, 16071615,  DOI: 10.1016/j.str.2014.09.007
  50. 50
    Kuo, A.; Bowler, M. W.; Zimmer, J.; Antcliff, J. F.; Doyle, D. A. Increasing the diffraction limit and internal order of a membrane protein crystal by dehydration. J. Struct. Biol. 2003, 141, 97102,  DOI: 10.1016/S1047-8477(02)00633-0
  51. 51
    Russo Krauss, I.; Sica, F.; Mattia, C. A.; Merlino, A. Increasing the X-ray Diffraction Power of Protein Crystals by Dehydration: The Case of Bovine Serum Albumin and a Survey of Literature Data. Int. J. Mol. Sci. 2012, 13, 37823800,  DOI: 10.3390/ijms13033782
  52. 52
    Kwan, T. O. C.; Axford, D.; Moraes, I. Membrane protein crystallography in the era of modern structural biology. Biochem. Soc. Trans. 2020, 48, 25052524,  DOI: 10.1042/BST20200066
  53. 53
    Müh, F.; Zouni, A. Structural basis of light-harvesting in the photosystem II core complex. Protein Sci. 2020, 29, 10901119,  DOI: 10.1002/pro.3841
  54. 54
    Birch, J.; Axford, D.; Foadi, J.; Meyer, A.; Eckhardt, A.; Thielmann, Y.; Moraes, I. The fine art of integral membrane protein crystallisation. Methods 2018, 147, 150162,  DOI: 10.1016/j.ymeth.2018.05.014
  55. 55
    Park, H.; Tran, T.; Lee, J. H.; Park, H.; Disney, M. D. Controlled dehydration improves the diffraction quality of two RNA crystals. BMC Struct. Biol. 2016, 16, 19,  DOI: 10.1186/s12900-016-0069-1
  56. 56
    Francia, F.; Palazzo, G.; Mallardi, A.; Cordone, L.; Venturoli, G. Residual water modulates QA-to-QB electron transfer in bacterial reaction centers embedded in trehalose amorphous matrices. Biophys. J. 2003, 85, 27602775,  DOI: 10.1016/S0006-3495(03)74698-0
  57. 57
    Palazzo, G.; Francia, F.; Mallardi, A.; Giustini, M.; Lopez, F.; Venturoli, G. Water activity regulates the QA to QB electron transfer in photosynthetic reaction centers from Rhodobacter sphaeroides. J. Am. Chem. Soc. 2008, 130, 93539363,  DOI: 10.1021/ja801963a
  58. 58
    Zabelin, A. A.; Khristin, A. M.; Shkuropatova, V. A.; Khatypov, R. A.; Shkuropatov, A. Y. Primary electron transfer in Rhodobacter sphaeroides R-26 reaction centers under dehydration conditions. Biochim. Biophys. Acta, Bioenerg. 2020, 1861, 148238  DOI: 10.1016/j.bbabio.2020.148238
  59. 59
    Noguchi, T.; Sugiura, M. Flash-Induced FTIR Difference Spectra of the Water Oxidizing Complex in Moderately Hydrated Photosystem II Core Films: Effect of Hydration Extent on S-State Transitions. Biochemistry 2002, 41, 23222330,  DOI: 10.1021/bi011954k
  60. 60
    Kaminskaya, O.; Renger, G.; Shuvalov, V. A. Effect of Dehydration on Light-Induced Reactions in Photosystem II: Photoreactions of Cytochrome b559. Biochemistry 2003, 42, 81198132,  DOI: 10.1021/bi020606v
  61. 61
    Pieper, J.; Hauss, T.; Buchsteiner, A.; Baczyński, K.; Adamiak, K.; Lechner, R. E.; Renger, G. Temperature- and Hydration-Dependent Protein Dynamics in Photosystem II of Green Plants Studied by Quasielastic Neutron Scattering. Biochemistry 2007, 46, 1139811409,  DOI: 10.1021/bi700179s
  62. 62
    Beglov, D.; Roux, B. An Integral Equation To Describe the Solvation of Polar Molecules in Liquid Water. J. Phys. Chem. B 1997, 101, 78217826,  DOI: 10.1021/jp971083h
  63. 63
    Kovalenko, A.; Hirata, F. Potential of mean force between two molecular ions in a polar molecular solvent: A study by the three-dimensional reference interaction site model. J. Phys. Chem. B 1999, 103, 79427957,  DOI: 10.1021/jp991300+
  64. 64
    Ben-Shalom, I. Y.; Lin, C.; Kurtzman, T.; Walker, R. C.; Gilson, M. K. Simulating Water Exchange to Buried Binding Sites. J. Chem. Theory Comput. 2019, 15, 26842691,  DOI: 10.1021/acs.jctc.8b01284
  65. 65
    Sindhikara, D. J.; Yoshida, N.; Hirata, F. Placevent: An algorithm for prediction of explicit solvent atom distribution─Application to HIV-1 protease and F-ATP synthase. J. Comput. Chem. 2012, 33, 15361543,  DOI: 10.1002/jcc.22984
  66. 66
    Koua, F. H. M.; Umena, Y.; Kawakami, K.; Shen, J.-R. Structure of Sr-Substituted Photosystem II at 2.1 Å Resolution and its Implications in the Mechanism of Water Oxidation. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 38893894,  DOI: 10.1073/pnas.1219922110
  67. 67
    Kato, K.; Miyazaki, N.; Hamaguchi, T.; Nakajima, Y.; Akita, F.; Yonekura, K.; Shen, J.-R. High-resolution cryo-EM structure of photosystem II reveals damage from high-dose electron beams. Commun. Biol. 2021, 4, 382,  DOI: 10.1038/s42003-021-01919-3
  68. 68
    Gisriel, C. J.; Wang, J.; Liu, J.; Flesher, D. A.; Reiss, K. M.; Huang, H.-L.; Yang, K. R.; Armstrong, W. H.; Gunner, M. R.; Batista, V. S.; Debus, R. J.; Brudvig, G. W. High-resolution cryo-electron microscopy structure of photosystem II from the mesophilic cyanobacterium, Synechocystis sp. PCC 6803. Proc. Natl. Acad. Sci. U. S. A. 2022, 119, e2116765118  DOI: 10.1073/pnas.2116765118
  69. 69
    Ogata, K.; Yuki, T.; Hatakeyama, M.; Uchida, W.; Nakamura, S. All-Atom Molecular Dynamics Simulation of Photosystem II Embedded in Thylakoid Membrane. J. Am. Chem. Soc. 2013, 135, 1567015673,  DOI: 10.1021/ja404317d
  70. 70
    Chrysina, M.; de Mendonça Silva, J. C.; Zahariou, G.; Pantazis, D. A.; Ioannidis, N. Proton Translocation via Tautomerization of Asn298 During the S2–S3 State Transition in the Oxygen-Evolving Complex of Photosystem II. J. Phys. Chem. B 2019, 123, 30683078,  DOI: 10.1021/acs.jpcb.9b02317
  71. 71
    Nakamura, S.; Nagao, R.; Takahashi, R.; Noguchi, T. Fourier Transform Infrared Detection of a Polarizable Proton Trapped between Photooxidized Tyrosine YZ and a Coupled Histidine in Photosystem II: Relevance to the Proton Transfer Mechanism of Water Oxidation. Biochemistry 2014, 53, 31313144,  DOI: 10.1021/bi500237y
  72. 72
    Kawashima, K.; Saito, K.; Ishikita, H. Mechanism of Radical Formation in the H-Bond Network of D1-Asn298 in Photosystem II. Biochemistry 2018, 57, 49975004,  DOI: 10.1021/acs.biochem.8b00574
  73. 73
    Xiao, Y.; Zhu, Q.; Yang, Y.; Wang, W.; Kuang, T.; Shen, J.-R.; Han, G. Role of PsbV-Tyr137 in photosystem II studied by site-directed mutagenesis in the thermophilic cyanobacterium Thermosynechococcus vulcanus. Photosynth. Res. 2020, 146, 4154,  DOI: 10.1007/s11120-020-00753-8
  74. 74
    Zhu, Q.; Yang, Y.; Xiao, Y.; Han, W.; Li, X.; Wang, W.; Kuang, T.; Shen, J. R.; Han, G. Effects of mutations of D1-R323, D1-N322, D1-D319, D1-H304 on the functioning of photosystem II in Thermosynechococcus vulcanus. Photosynth. Res. 2022, 152, 193206,  DOI: 10.1007/s11120-022-00920-z
  75. 75
    Müh, F.; Plöckinger, M.; Renger, T. Electrostatic Asymmetry in the Reaction Center of Photosystem II. J. Phys. Chem. Lett. 2017, 8, 850858,  DOI: 10.1021/acs.jpclett.6b02823
  76. 76
    Sirohiwal, A.; Neese, F.; Pantazis, D. A. Protein Matrix Control of Reaction Center Excitation in Photosystem II. J. Am. Chem. Soc. 2020, 142, 1817418190,  DOI: 10.1021/jacs.0c08526
  77. 77
    Holzwarth, A. R.; Müller, M. G.; Reus, M.; Nowaczyk, M.; Sander, J.; Rögner, M. Kinetics and mechanism of electron transfer in intact photosystem II and in the isolated reaction center: Pheophytin is the primary electron acceptor. Proc. Natl. Acad. Sci. U. S. A. 2006, 103, 68956900,  DOI: 10.1073/pnas.0505371103
  78. 78
    Prokhorenko, V. I.; Holzwarth, A. R. Primary Processes and Structure of the Photosystem II Reaction Center: A Photon Echo Study. J. Phys. Chem. B 2000, 104, 1156311578,  DOI: 10.1021/jp002323n
  79. 79
    Saito, K.; William Rutherford, A.; Ishikita, H. Energetics of proton release on the first oxidation step in the water-oxidizing enzyme. Nat. Commun. 2015, 6, 8488,  DOI: 10.1038/ncomms9488
  80. 80
    Reiss, K.; Morzan, U. N.; Grigas, A. T.; Batista, V. S. Water Network Dynamics Next to the Oxygen-Evolving Complex of Photosystem II. Inorganics 2019, 7, 39,  DOI: 10.3390/inorganics7030039
  81. 81
    Pal, R.; Negre, C. F. A.; Vogt, L.; Pokhrel, R.; Ertem, M. Z.; Brudvig, G. W.; Batista, V. S. S0-state model of the oxygen-evolving complex of photosystem II. Biochemistry 2013, 52, 77037706,  DOI: 10.1021/bi401214v
  82. 82
    Krewald, V.; Retegan, M.; Cox, N.; Messinger, J.; Lubitz, W.; DeBeer, S.; Neese, F.; Pantazis, D. A. Metal Oxidation States in Biological Water Splitting. Chem. Sci. 2015, 6, 16761695,  DOI: 10.1039/C4SC03720K
  83. 83
    Lohmiller, T.; Krewald, V.; Sedoud, A.; Rutherford, A. W.; Neese, F.; Lubitz, W.; Pantazis, D. A.; Cox, N. The First State in the Catalytic Cycle of the Water-Oxidizing Enzyme: Identification of a Water-Derived μ-Hydroxo Bridge. J. Am. Chem. Soc. 2017, 139, 1441214424,  DOI: 10.1021/jacs.7b05263
  84. 84
    Shimada, Y.; Sugiyama, A.; Nagao, R.; Noguchi, T. Role of D1-Glu65 in Proton Transfer during Photosynthetic Water Oxidation in Photosystem II. J. Phys. Chem. B 2022, 126, 82028213,  DOI: 10.1021/acs.jpcb.2c05869
  85. 85
    Allgöwer, F.; Gamiz-Hernandez, A. P.; Rutherford, A. W.; Kaila, V. R. I. Molecular Principles of Redox-Coupled Protonation Dynamics in Photosystem II. J. Am. Chem. Soc. 2022, 144, 71717180,  DOI: 10.1021/jacs.1c13041
  86. 86
    Pokhrel, R.; Service, R. J.; Debus, R. J.; Brudvig, G. W. Mutation of Lysine 317 in the D2 Subunit of Photosystem II Alters Chloride Binding and Proton Transport. Biochemistry 2013, 52, 47584773,  DOI: 10.1021/bi301700u
  87. 87
    Rivalta, I.; Amin, M.; Luber, S.; Vassiliev, S.; Pokhrel, R.; Umena, Y.; Kawakami, K.; Shen, J. R.; Kamiya, N.; Bruce, D.; Brudvig, G. W.; Gunner, M. R.; Batista, V. S. Structural-Functional Role of Chloride in Photosystem II. Biochemistry 2011, 50, 63126315,  DOI: 10.1021/bi200685w
  88. 88
    Suzuki, H.; Yu, J.; Kobayashi, T.; Nakanishi, H.; Nixon, P. J.; Noguchi, T. Functional Roles of D2-Lys317 and the Interacting Chloride Ion in the Water Oxidation Reaction of Photosystem II As Revealed by Fourier Transform Infrared Analysis. Biochemistry 2013, 52, 47484757,  DOI: 10.1021/bi301699h
  89. 89
    Kuroda, H.; Kawashima, K.; Ueda, K.; Ikeda, T.; Saito, K.; Ninomiya, R.; Hida, C.; Takahashi, Y.; Ishikita, H. Proton transfer pathway from the oxygen-evolving complex in photosystem II substantiated by extensive mutagenesis. Biochim. Biophys. Acta, Bioenerg. 2021, 1862, 148329  DOI: 10.1016/j.bbabio.2020.148329
  90. 90
    Kaur, D.; Zhang, Y.; Reiss, K. M.; Mandal, M.; Brudvig, G. W.; Batista, V. S.; Gunner, M. R. Proton exit pathways surrounding the oxygen evolving complex of photosystem II. Biochim. Biophys. Acta, Bioenerg. 2021, 1862, 148446  DOI: 10.1016/j.bbabio.2021.148446
  91. 91
    Isobe, H.; Shoji, M.; Suzuki, T.; Shen, J.-R.; Yamaguchi, K. Spin, Valence, and Structural Isomerism in the S3 State of the Oxygen-Evolving Complex of Photosystem II as a Manifestation of Multimetallic Cooperativity. J. Chem. Theory Comput. 2019, 15, 23752391,  DOI: 10.1021/acs.jctc.8b01055
  92. 92
    Pantazis, D. A. The S3 State of the Oxygen-Evolving Complex: Overview of Spectroscopy and XFEL Crystallography with a Critical Evaluation of Early-Onset Models for O–O Bond Formation. Inorganics 2019, 7, 55,  DOI: 10.3390/inorganics7040055
  93. 93
    Corry, T. A.; O’Malley, P. J. S3 State Models of Nature’s Water Oxidizing Complex: Analysis of Bonding and Magnetic Exchange Pathways, Assessment of Experimental Electron Paramagnetic Resonance Data, and Implications for the Water Oxidation Mechanism. J. Phys. Chem. B 2021, 125, 1009710107,  DOI: 10.1021/acs.jpcb.1c04459
  94. 94
    Drosou, M.; Pantazis, D. A. Redox Isomerism in the S3 State of the Oxygen-Evolving Complex Resolved by Coupled Cluster Theory. Chem. – Eur. J. 2021, 27, 1281512825,  DOI: 10.1002/chem.202101567
  95. 95
    Okamoto, Y.; Shimada, Y.; Nagao, R.; Noguchi, T. Proton and Water Transfer Pathways in the S2 → S3 Transition of the Water-Oxidizing Complex in Photosystem II: Time-Resolved Infrared Analysis of the Effects of D1-N298A Mutation and NO3 Substitution. J. Phys. Chem. B 2021, 125, 68646873,  DOI: 10.1021/acs.jpcb.1c03386
  96. 96
    Oyala, P. H.; Stich, T. A.; Stull, J. A.; Yu, F.; Pecoraro, V. L.; Britt, R. D. Pulse Electron Paramagnetic Resonance Studies of the Interaction of Methanol with the S2 State of the Mn4O5Ca Cluster of Photosystem II. Biochemistry 2014, 53, 79147928,  DOI: 10.1021/bi501323h
  97. 97
    Kalendra, V.; Reiss, K. M.; Banerjee, G.; Ghosh, I.; Baldansuren, A.; Batista, V. S.; Brudvig, G. W.; Lakshmi, K. V. Binding of the substrate analog methanol in the oxygen-evolving complex of photosystem II in the D1-N87A genetic variant of cyanobacteria. Faraday Discuss. 2022, 234, 195213,  DOI: 10.1039/D1FD00094B
  98. 98
    Pérez Navarro, M.; Ames, W. M.; Nilsson, H.; Lohmiller, T.; Pantazis, D. A.; Rapatskiy, L.; Nowaczyk, M. M.; Neese, F.; Boussac, A.; Messinger, J.; Lubitz, W.; Cox, N. Ammonia binding to the oxygen-evolving complex of photosystem II identifies the solvent-exchangeable oxygen bridge (μ-oxo) of the manganese tetramer. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 1556115566,  DOI: 10.1073/pnas.1304334110
  99. 99
    Schraut, J.; Kaupp, M. On Ammonia Binding to the Oxygen-Evolving Complex of Photosystem II: A Quantum Chemical Study. Chem. – Eur. J. 2014, 20, 73007308,  DOI: 10.1002/chem.201304464
  100. 100
    Oyala, P. H.; Stich, T. A.; Debus, R. J.; Britt, R. D. Ammonia Binds to the Dangler Manganese of the Photosystem II Oxygen Evolving Complex. J. Am. Chem. Soc. 2015, 137, 88298837,  DOI: 10.1021/jacs.5b04768
  101. 101
    Guo, Y.; He, L.-L.; Zhao, D.-X.; Gong, L.-D.; Liu, C.; Yang, Z.-Z. How does ammonia bind to the oxygen-evolving complex in the S2 state of photosynthetic water oxidation? Theoretical support and implications for the W1 substitution mechanism. Phys. Chem. Chem. Phys. 2016, 18, 3155131565,  DOI: 10.1039/C6CP05725J
  102. 102
    Marchiori, D. A.; Oyala, P. H.; Debus, R. J.; Stich, T. A.; Britt, R. D. Structural Effects of Ammonia Binding to the Mn4CaO5 Cluster of Photosystem II. J. Phys. Chem. B 2018, 122, 15881599,  DOI: 10.1021/acs.jpcb.7b11101
  103. 103
    Askerka, M.; Vinyard, D. J.; Brudvig, G. W.; Batista, V. S. NH3 Binding to the S2 State of the O2-Evolving Complex of Photosystem II: Analogue to H2O Binding during the S2 → S3 Transition. Biochemistry 2015, 54, 57835786,  DOI: 10.1021/acs.biochem.5b00974
  104. 104
    Schuth, N.; Liang, Z.; Schönborn, M.; Kussicke, A.; Assunção, R.; Zaharieva, I.; Zilliges, Y.; Dau, H. Inhibitory and Non-Inhibitory NH3 Binding at the Water-Oxidizing Manganese Complex of Photosystem II Suggests Possible Sites and a Rearrangement Mode of Substrate Water Molecules. Biochemistry 2017, 56, 62406256,  DOI: 10.1021/acs.biochem.7b00743
  105. 105
    Shevela, D.; Eaton-Rye, J. J.; Shen, J.-R.; Govindjee Photosystem II and the unique role of bicarbonate: a historical perspective. Biochim. Biophys. Acta, Bioenerg. 2012, 1817, 11341151,  DOI: 10.1016/j.bbabio.2012.04.003
  106. 106
    Müh, F.; Glöckner, C.; Hellmich, J.; Zouni, A. Light-induced quinone reduction in photosystem II. Biochim. Biophys. Acta, Bioenerg. 2012, 1817, 4465,  DOI: 10.1016/j.bbabio.2011.05.021
  107. 107
    Hutchison, R. S.; Xiong, J.; Sayre, R. T.; Govindjee Construction and characterization of a Photosystem II D1 mutant (arginine-269-glycine) of Chlamydomonas reinhardtii. Biochim. Biophys. Acta, Bioenerg. 1996, 1277, 8392,  DOI: 10.1016/S0005-2728(96)00085-0
  108. 108
    Petrouleas, V.; Crofts, A. R. In Photosystem II: The Light-Driven Water:Plastoquinone Oxidoreductase; Wydrzynski, T. J.; Satoh, K.; Freeman, J. A., Eds.; Springer Netherlands: Dordrecht, 2005, p 177206.
  109. 109
    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, 954959,  DOI: 10.1073/pnas.1212957110
  110. 110
    Forsman, J. A.; Eaton-Rye, J. J. The D1:Ser268 residue of Photosystem II contributes to an alternative pathway for QB protonation in the absence of bound bicarbonate. FEBS Lett. 2020, 594, 29532964,  DOI: 10.1002/1873-3468.13880
  111. 111
    Alfonso, M.; Pueyo, J. J.; Gaddour, K.; Etienne, A. L.; Kirilovsky, D.; Picorel, R. Induced New Mutation of D1 Serine-268 in Soybean Photosynthetic Cell Cultures Produced Atrazine Resistance, Increased Stability of S2QB and S3QB States, and Increased Sensitivity to Light Stress. Plant Physiol. 1996, 112, 14991508,  DOI: 10.1104/pp.112.4.1499
  112. 112
    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, Bioenerg. 2019, 1860, 148054,  DOI: 10.1016/j.bbabio.2019.07.009
  113. 113
    Khaing, E. P.; Zhong, V.; Kodru, S.; Vass, I.; Eaton-Rye, J. J. Tyr244 of the D2 Protein Is Required for Correct Assembly and Operation of the Quinone-Iron-Bicarbonate Acceptor Complex of Photosystem II. Biochemistry 2022, 61, 12981312,  DOI: 10.1021/acs.biochem.2c00164
  114. 114
    Berthomieu, C.; Hienerwadel, R. Iron coordination in photosystem II: interaction between bicarbonate and the QB pocket studied by Fourier transform infrared spectroscopy. Biochemistry 2001, 40, 40444052,  DOI: 10.1021/bi002236l
  115. 115
    Takahashi, R.; Boussac, A.; Sugiura, M.; Noguchi, T. Structural coupling of a tyrosine side chain with the non-heme iron center in photosystem II as revealed by light-induced Fourier transform infrared difference spectroscopy. Biochemistry 2009, 48, 89949001,  DOI: 10.1021/bi901195e
  116. 116
    Cardona, T.; Sedoud, A.; Cox, N.; Rutherford, A. W. Charge Separation in Photosystem II: A Comparative and Evolutionary Overview. Biochim. Biophys. Acta, Bioenerg. 2012, 1817, 2643,  DOI: 10.1016/j.bbabio.2011.07.012
  117. 117
    Okamura, M. Y.; Paddock, M. L.; Graige, M. S.; Feher, G. Proton and electron transfer in bacterial reaction centers. Biochim. Biophys. Acta, Bioenerg. 2000, 1458, 148163,  DOI: 10.1016/S0005-2728(00)00065-7
  118. 118
    de Wijn, R.; van Gorkom, H. J. Kinetics of Electron Transfer from QA to QB in Photosystem II. Biochemistry 2001, 40, 1191211922,  DOI: 10.1021/bi010852r
  119. 119
    Paddock, M. L.; Ädelroth, P.; Chang, C.; Abresch, E. C.; Feher, G.; Okamura, M. Y. Identification of the Proton Pathway in Bacterial Reaction Centers: Cooperation between Asp-M17 and Asp-L210 Facilitates Proton Transfer to the Secondary Quinone (QB). Biochemistry 2001, 40, 68936902,  DOI: 10.1021/bi010280a
  120. 120
    Paddock, M. L.; Feher, G.; Okamura, M. Y. Proton transfer pathways and mechanism in bacterial reaction centers. FEBS Lett. 2003, 555, 4550,  DOI: 10.1016/S0014-5793(03)01149-9
  121. 121
    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  DOI: 10.1073/pnas.2103203118
  122. 122
    Fantuzzi, A.; Allgöwer, F.; Baker, H.; McGuire, G.; Teh, W. K.; Gamiz-Hernandez, A. P.; Kaila, V. R. I.; Rutherford, A. W. Bicarbonate-controlled reduction of oxygen by the QA semiquinone in Photosystem II in membranes. Proc. Natl. Acad. Sci. U. S. A. 2022, 119, e2116063119  DOI: 10.1073/pnas.2116063119
  123. 123
    Sugo, Y.; Saito, K.; Ishikita, H. Conformational Changes and H-Bond Rearrangements during Quinone Release in Photosystem II. Biochemistry 2022, 61, 18361843,  DOI: 10.1021/acs.biochem.2c00324

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  2. Richard J. Debus, Paul H. Oyala. Independent Mutation of Two Bridging Carboxylate Ligands Stabilizes Alternate Conformers of the Photosynthetic O2-Evolving Mn4CaO5 Cluster in Photosystem II. The Journal of Physical Chemistry B 2024, Article ASAP.
  3. Takumi Noguchi. Mechanism of Proton Transfer through the D1-E65/D2-E312 Gate during Photosynthetic Water Oxidation. The Journal of Physical Chemistry B 2024, 128 (8) , 1866-1875. https://doi.org/10.1021/acs.jpcb.3c07787
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  5. Abhishek Sirohiwal, Dimitrios A. Pantazis. Reaction Center Excitation in Photosystem II: From Multiscale Modeling to Functional Principles. Accounts of Chemical Research 2023, 56 (21) , 2921-2932. https://doi.org/10.1021/acs.accounts.3c00392
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  • Abstract

    Figure 1

    Figure 1. (a) Model of the membrane-embedded PSII monomer. Key protein subunits are labeled. The location of the oxygen-evolving complex and the non-heme iron are indicated. The PsbO, PsbU, and PsbV proteins cap cyanobacterial PSII on the lumenal side of the membrane. (b) Redox-active cofactors in the core proteins of PSII and the two reactions accomplished at the oxidative and reductive termini. The arrows indicate the normal flow of electrons.

    Figure 2

    Figure 2. (a) Flow-chart describing the simulation procedure employed in the current work. (b) Time evolution of root mean square deviation (RMSD) of Cα atoms of the PSII monomer during production simulations (data recorded every 2 ps, excluding the highly flexible loops). (c) Time evolution of the hydration content of the PSII monomer. All water molecules in close contact (∼2 Å) with PSII were selected in the counting (data recorded every 50 ps, hydration content around the highly flexible loops not considered). (d) Time evolution of water content within a sphere of radius 20 Å centered around the OEC (data recorded every 2 ps). Corresponding datasets that include the water content associated with the highly flexible loops are provided in the Supporting Information.

    Figure 3

    Figure 3. Identification of water channels leading from the bulk toward the OEC, along with the proposal for the proton exit pathway associated with YZ. O1 (blue), O4 (red), and Cl1 (green) channel systems are colored uniquely for presentation purposes. Asterisks indicate positions of DMSO molecules found in the 5B66 crystallographic model. Dotted lines imply the transient nature of respective water channels due to protein conformational dynamics.

    Figure 4

    Figure 4. Depiction of the E1 branch of the O1 channel system.

    Figure 5

    Figure 5. Depiction of the E2 branch of the O1 channel system. The superscript CTR denotes C-terminal residues.

    Figure 6

    Figure 6. Depiction of the E3 and E4 branches of the O1 channel system, along with the YZ water network that converges to E4 close to the D1-Asp319 residue.

    Figure 7

    Figure 7. Role of the PsbV-Tyr137 residue in gating the E3 and E4 channels by adopting two distinct orientations.

    Figure 8

    Figure 8. Depiction of the E5, E6, and E7 branches of the Cl1 channel system.

    Figure 9

    Figure 9. Transient water wire formation that is dependent on the conformation of D1-Val67: open (left) and closed (right) configurations determined by the side-chain rotation of D1-Val67.

    Figure 10

    Figure 10. Depiction of the O4 channel system. The E8 and E9 branches merge close to the PsbU-Asp96/CP43-Lys339 pair.

    Figure 11

    Figure 11. Schematic figure showing the three water channel systems connecting the non-heme iron site of PSII with the stromal bulk. Asterisks indicate positions of DMSO molecules found in the 5B66 crystallographic model. Dotted lines imply the transient nature of respective water channels due to protein conformational dynamics.

    Figure 12

    Figure 12. (a) Identification of the A1 water network leading from the bulk to the non-heme iron site; (b) A2 channel in the closed state, channel A1 is also depicted; (c) A2 channel in the open state, merging with channel A1.

    Figure 13

    Figure 13. B1 (left) and B2 (right) water branches connecting the bulk with the NHI site.

    Figure 14

    Figure 14. (a) Entry of water into channel C blocked by the hydrophobic side chains of D1-Ile248 and D1-Leu271; (b) formation of water wire connecting the bicarbonate site through the QB pocket with the bulk; (c) QB pocket water trapped because exit is restricted by the side chain conformation of D1-Asn266.

    Figure 15

    Figure 15. Differences in the protein environment around the analogous quinones in (a) the Photosystem II (PDB ID 3WU2) and (b) the bacterial reaction center (PDB ID 3I4D).

  • References

    ARTICLE SECTIONS
    Jump To

    This article references 123 other publications.

    1. 1
      Shevela, D.; Björn, L. O.; Govindjee. Photosynthesis: Solar Energy for Life; World Scientific: Singapore, 2017, p 204.
    2. 2
      Blankenship, R. E. Molecular Mechanisms of Photosynthesis; 2nd ed.; Wiley: Chichester, 2014, p 312.
    3. 3
      Photosystem II. The Light-Driven Water:Plastoquinone Oxidoreductase; Wydrzynski, T.; Satoh, K., Eds.; Springer: Dordrecht, 2005; Vol. 22, p 786.
    4. 4
      Barber, J. Photosystem II: the water splitting enzyme of photosynthesis and the origin of oxygen in our atmosphere. Q. Rev. Biophys. 2016, 49, e14  DOI: 10.1017/S0033583516000093
    5. 5
      Krewald, V.; Retegan, M.; Pantazis, D. A. Principles of Natural Photosynthesis. Top. Curr. Chem. 2016, 371, 2348,  DOI: 10.1007/128_2015_645
    6. 6
      Shen, J.-R. The Structure of Photosystem II and the Mechanism of Water Oxidation in Photosynthesis. Annu. Rev. Plant Biol. 2015, 66, 2348,  DOI: 10.1146/annurev-arplant-050312-120129
    7. 7
      Pantazis, D. A. In Hydrogen Production and Energy Transition; Van de Voorde, M., Ed.; De Gruyter: 2021; Vol. 1, p 427468.
    8. 8
      Junge, W. Oxygenic photosynthesis: history, status and perspective. Q. Rev. Biophys. 2019, 52, e1  DOI: 10.1017/S0033583518000112
    9. 9
      Pantazis, D. A. Missing Pieces in the Puzzle of Biological Water Oxidation. ACS Catal. 2018, 8, 94779507,  DOI: 10.1021/acscatal.8b01928
    10. 10
      Cox, N.; Pantazis, D. A.; Lubitz, W. Current Understanding of the Mechanism of Water Oxidation in Photosystem II and Its Relation to XFEL Data. Annu. Rev. Biochem. 2020, 89, 795820,  DOI: 10.1146/annurev-biochem-011520-104801
    11. 11
      Ho, F. M. Uncovering channels in photosystem II by computer modelling: current progress, future prospects, and lessons from analogous systems. Photosynth. Res. 2008, 98, 503522,  DOI: 10.1007/s11120-008-9358-2
    12. 12
      Ho, F. M.; Styring, S. Access Channels and Methanol Binding Site to the CaMn4 Cluster in Photosystem II Based on Solvent Accessibility Simulations, with Implications for Substrate Water Access. Biochim. Biophys. Acta, Bioenerg. 2008, 1777, 140153,  DOI: 10.1016/j.bbabio.2007.08.009
    13. 13
      Ho, F. M. In Molecular Solar Fuels; Wydrzynski, T. J.; Hillier, W., Eds.; The Royal Society of Chemistry: Cambridge, 2012, p 208248.
    14. 14
      Linke, K.; Ho, F. M. Water in Photosystem II: Structural, Functional and Mechanistic Considerations. Biochim. Biophys. Acta, Bioenerg. 2014, 1837, 1432,  DOI: 10.1016/j.bbabio.2013.08.003
    15. 15
      Nakamura, S.; Ota, K.; Shibuya, Y.; Noguchi, T. Role of a Water Network around the Mn4CaO5 Cluster in Photosynthetic Water Oxidation: A Fourier Transform Infrared Spectroscopy and Quantum Mechanics/Molecular Mechanics Calculation Study. Biochemistry 2016, 55, 597607,  DOI: 10.1021/acs.biochem.5b01120
    16. 16
      Bondar, A.-N.; Dau, H. Extended Protein/Water H-bond Networks in Photosynthetic Water Oxidation. Biochim. Biophys. Acta, Bioenerg. 2012, 1817, 11771190,  DOI: 10.1016/j.bbabio.2012.03.031
    17. 17
      Murray, J.; Barber, J. In Photosynthesis. Energy from the Sun; Allen, J.; Gantt, E.; Golbeck, J.; Osmond, B., Eds.; Springer Netherlands: 2008, p 467470,  DOI: 10.1007/978-1-4020-6709-9_105 .
    18. 18
      Gabdulkhakov, A.; Guskov, A.; Broser, M.; Kern, J.; Müh, F.; Saenger, W.; Zouni, A. Probing the Accessibility of the Mn4Ca Cluster in Photosystem II: Channels Calculation, Noble Gas Derivatization, and Cocrystallization with DMSO. Structure 2009, 17, 12231234,  DOI: 10.1016/j.str.2009.07.010
    19. 19
      Vassiliev, S.; Comte, P.; Mahboob, A.; Bruce, D. Tracking the flow of water through photosystem II using molecular dynamics and streamline tracing. Biochemistry 2010, 49, 18731881,  DOI: 10.1021/bi901900s
    20. 20
      Vassiliev, S.; Zaraiskaya, T.; Bruce, D. Exploring the Energetics of Water Permeation in Photosystem II by Multiple Steered Molecular Dynamics Simulations. Biochim. Biophys. Acta, Bioenerg. 2012, 1817, 16711678,  DOI: 10.1016/j.bbabio.2012.05.016
    21. 21
      Vassiliev, S.; Zaraiskaya, T.; Bruce, D. Molecular Dynamics Simulations Reveal Highly Permeable Oxygen Exit Channels Shared with Water Uptake Channels in Photosystem II. Biochim. Biophys. Acta, Bioenerg. 2013, 1827, 11481155,  DOI: 10.1016/j.bbabio.2013.06.008
    22. 22
      Ishikita, H.; Saenger, W.; Loll, B.; Biesiadka, J.; Knapp, E.-W. Energetics of a Possible Proton Exit Pathway for Water Oxidation in Photosystem II. Biochemistry 2006, 45, 20632071,  DOI: 10.1021/bi051615h
    23. 23
      Takaoka, T.; Sakashita, N.; Saito, K.; Ishikita, H. pKa of a Proton-Conducting Water Chain in Photosystem II. J. Phys. Chem. Lett. 2016, 7, 19251932,  DOI: 10.1021/acs.jpclett.6b00656
    24. 24
      Sakashita, N.; Watanabe, H. C.; Ikeda, T.; Ishikita, H. Structurally conserved channels in cyanobacterial and plant photosystem II. Photosynth. Res. 2017, 133, 7585,  DOI: 10.1007/s11120-017-0347-1
    25. 25
      Sakashita, N.; Watanabe, H. C.; Ikeda, T.; Saito, K.; Ishikita, H. Origins of Water Molecules in the Photosystem II Crystal Structure. Biochemistry 2017, 56, 30493057,  DOI: 10.1021/acs.biochem.7b00220
    26. 26
      Sakashita, N.; Ishikita, H.; Saito, K. Rigidly hydrogen-bonded water molecules facilitate proton transfer in photosystem II. Phys. Chem. Chem. Phys. 2020, 22, 1583115841,  DOI: 10.1039/D0CP00295J
    27. 27
      Ogata, K.; Hatakeyama, M.; Sakamoto, Y.; Nakamura, S. Investigation of a Pathway for Water Delivery in Photosystem II Protein by Molecular Dynamics Simulation. J. Phys. Chem. B 2019, 123, 64446452,  DOI: 10.1021/acs.jpcb.9b04838
    28. 28
      Ghosh, I.; Khan, S.; Banerjee, G.; Dziarski, A.; Vinyard, D. J.; Debus, R. J.; Brudvig, G. W. Insights into Proton-Transfer Pathways during Water Oxidation in Photosystem II. J. Phys. Chem. B 2019, 123, 81958202,  DOI: 10.1021/acs.jpcb.9b06244
    29. 29
      Kaur, D.; Cai, X.; Khaniya, U.; Zhang, Y.; Mao, J.; Mandal, M.; Gunner, M. R. Tracing the Pathways of Waters and Protons in Photosystem II and Cytochrome c Oxidase. Inorganics 2019, 7, 14,  DOI: 10.3390/inorganics7020014
    30. 30
      Guerra, F.; Siemers, M.; Mielack, C.; Bondar, A.-N. Dynamics of Long-Distance Hydrogen-Bond Networks in Photosystem II. J. Phys. Chem. B 2018, 122, 46254641,  DOI: 10.1021/acs.jpcb.8b00649
    31. 31
      Saito, K.; Rutherford, A. W.; Ishikita, H. Mechanism of Tyrosine D Oxidation in Photosystem II. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 76907695,  DOI: 10.1073/pnas.1300817110
    32. 32
      Sirohiwal, A.; Neese, F.; Pantazis, D. A. Microsolvation of the Redox-Active Tyrosine-D in Photosystem II: Correlation of Energetics with EPR Spectroscopy and Oxidation-Induced Proton Transfer. J. Am. Chem. Soc. 2019, 141, 32173231,  DOI: 10.1021/jacs.8b13123
    33. 33
      Hussein, R.; Ibrahim, M.; Bhowmick, A.; Simon, P. S.; Chatterjee, R.; Lassalle, L.; Doyle, M.; Bogacz, I.; Kim, I.-S.; Cheah, M. H.; Gul, S.; de Lichtenberg, C.; Chernev, P.; Pham, C. C.; Young, I. D.; Carbajo, S.; Fuller, F. D.; Alonso-Mori, R.; Batyuk, A.; Sutherlin, K. D.; Brewster, A. S.; Bolotovsky, R.; Mendez, D.; Holton, J. M.; Moriarty, N. W.; Adams, P. D.; Bergmann, U.; Sauter, N. K.; Dobbek, H.; Messinger, J.; Zouni, A.; Kern, J.; Yachandra, V. K.; Yano, J. Structural dynamics in the water and proton channels of photosystem II during the S2 to S3 transition. Nat. Commun. 2021, 12, 6531,  DOI: 10.1038/s41467-021-26781-z
    34. 34
      Retegan, M.; Pantazis, D. A. Interaction of methanol with the oxygen-evolving complex: atomistic models, channel identification, species dependence, and mechanistic implications. Chem. Sci. 2016, 7, 64636476,  DOI: 10.1039/C6SC02340A
    35. 35
      Retegan, M.; Pantazis, D. A. Differences in the Active Site of Water Oxidation among Photosynthetic Organisms. J. Am. Chem. Soc. 2017, 139, 1434014343,  DOI: 10.1021/jacs.7b06351
    36. 36
      Vogt, L.; Vinyard, D. J.; Khan, S.; Brudvig, G. W. Oxygen-evolving complex of Photosystem II: an analysis of second-shell residues and hydrogen-bonding networks. Curr. Opin. Chem. Biol. 2015, 25, 152158,  DOI: 10.1016/j.cbpa.2014.12.040
    37. 37
      Weisz, D. A.; Gross, M. L.; Pakrasi, H. B. Reactive oxygen species leave a damage trail that reveals water channels in Photosystem II. Sci. Adv. 2017, 3, eaao3013  DOI: 10.1126/sciadv.aao3013
    38. 38
      Loll, B.; Kern, J.; Saenger, W.; Zouni, A.; Biesiadka, J. Towards complete cofactor arrangement in the 3.0 Å resolution structure of photosystem II. Nature 2005, 438, 10401044,  DOI: 10.1038/nature04224
    39. 39
      Ferreira, K. N.; Iverson, T. M.; Maghlaoui, K.; Barber, J.; Iwata, S. Architecture of the Photosynthetic Oxygen-Evolving Center. Science 2004, 303, 18311838,  DOI: 10.1126/science.1093087
    40. 40
      Murray, J. W.; Barber, J. Structural Characteristics of Channels and Pathways in Photosystem II Including the Identification of an Oxygen Channel. J. Struct. Biol. 2007, 159, 228237,  DOI: 10.1016/j.jsb.2007.01.016
    41. 41
      Umena, Y.; Kawakami, K.; Shen, J.-R.; Kamiya, N. Crystal Structure of the Oxygen-Evolving Photosystem II at a Resolution of 1.9 Å. Nature 2011, 473, 5560,  DOI: 10.1038/nature09913
    42. 42
      Suga, M.; Akita, F.; Hirata, K.; Ueno, G.; Murakami, H.; Nakajima, Y.; Shimizu, T.; Yamashita, K.; Yamamoto, M.; Ago, H.; Shen, J.-R. Native Structure of Photosystem II at 1.95 Å Resolution Viewed by Femtosecond X-ray Pulses. Nature 2015, 517, 99103,  DOI: 10.1038/nature13991
    43. 43
      Tanaka, A.; Fukushima, Y.; Kamiya, N. Two different structures of the oxygen-evolving complex in the same polypeptide frameworks of photosystem II. J. Am. Chem. Soc. 2017, 139, 17181721,  DOI: 10.1021/jacs.6b09666
    44. 44
      Suga, M.; Akita, F.; Sugahara, M.; Kubo, M.; Nakajima, Y.; Nakane, T.; Yamashita, K.; Umena, Y.; Nakabayashi, M.; Yamane, T.; Nakano, T.; Suzuki, M.; Masuda, T.; Inoue, S.; Kimura, T.; Nomura, T.; Yonekura, S.; Yu, L.-J.; Sakamoto, T.; Motomura, T.; Chen, J.-H.; Kato, Y.; Noguchi, T.; Tono, K.; Joti, Y.; Kameshima, T.; Hatsui, T.; Nango, E.; Tanaka, R.; Naitow, H.; Matsuura, Y.; Yamashita, A.; Yamamoto, M.; Nureki, O.; Yabashi, M.; Ishikawa, T.; Iwata, S.; Shen, J.-R. Light-Induced Structural Changes and the Site of O=O bond Formation in PSII Caught by XFEL. Nature 2017, 543, 131135,  DOI: 10.1038/nature21400
    45. 45
      Young, I. D.; Ibrahim, M.; Chatterjee, R.; Gul, S.; Fuller, F. D.; Koroidov, S.; Brewster, A. S.; Tran, R.; Alonso-Mori, R.; Kroll, T.; Michels-Clark, T.; Laksmono, H.; Sierra, R. G.; Stan, C. A.; Hussein, R.; Zhang, M.; Douthit, L.; Kubin, M.; de Lichtenberg, C.; Vo Pham, L.; Nilsson, H.; Cheah, M. H.; Shevela, D.; Saracini, C.; Bean, M. A.; Seuffert, I.; Sokaras, D.; Weng, T.-C.; Pastor, E.; Weninger, C.; Fransson, T.; Lassalle, L.; Bräuer, P.; Aller, P.; Docker, P. T.; Andi, B.; Orville, A. M.; Glownia, J. M.; Nelson, S.; Sikorski, M.; Zhu, D.; Hunter, M. S.; Lane, T. J.; Aquila, A.; Koglin, J. E.; Robinson, J.; Liang, M.; Boutet, S.; Lyubimov, A. Y.; Uervirojnangkoorn, M.; Moriarty, N. W.; Liebschner, D.; Afonine, P. V.; Waterman, D. G.; Evans, G.; Wernet, P.; Dobbek, H.; Weis, W. I.; Brunger, A. T.; Zwart, P. H.; Adams, P. D.; Zouni, A.; Messinger, J.; Bergmann, U.; Sauter, N. K.; Kern, J.; Yachandra, V. K.; Yano, J. Structure of Photosystem II and Substrate Binding at Room Temperature. Nature 2016, 540, 453457,  DOI: 10.1038/nature20161
    46. 46
      Kern, J.; Chatterjee, R.; Young, I. D.; Fuller, F. D.; Lassalle, L.; Ibrahim, M.; Gul, S.; Fransson, T.; Brewster, A. S.; Alonso-Mori, R.; Hussein, R.; Zhang, M.; Douthit, L.; de Lichtenberg, C.; Cheah, M. H.; Shevela, D.; Wersig, J.; Seuffert, I.; Sokaras, D.; Pastor, E.; Weninger, C.; Kroll, T.; Sierra, R. G.; Aller, P.; Butryn, A.; Orville, A. M.; Liang, M.; Batyuk, A.; Koglin, J. E.; Carbajo, S.; Boutet, S.; Moriarty, N. W.; Holton, J. M.; Dobbek, H.; Adams, P. D.; Bergmann, U.; Sauter, N. K.; Zouni, A.; Messinger, J.; Yano, J.; Yachandra, V. K. Structures of the Intermediates of Kok’s Photosynthetic Water Oxidation Clock. Nature 2018, 563, 421425,  DOI: 10.1038/s41586-018-0681-2
    47. 47
      Ibrahim, M.; Fransson, T.; Chatterjee, R.; Cheah, M. H.; Hussein, R.; Lassalle, L.; Sutherlin, K. D.; Young, I. D.; Fuller, F. D.; Gul, S.; Kim, I.-S.; Simon, P. S.; de Lichtenberg, C.; Chernev, P.; Bogacz, I.; Pham, C. C.; Orville, A. M.; Saichek, N.; Northen, T.; Batyuk, A.; Carbajo, S.; Alonso-Mori, R.; Tono, K.; Owada, S.; Bhowmick, A.; Bolotovsky, R.; Mendez, D.; Moriarty, N. W.; Holton, J. M.; Dobbek, H.; Brewster, A. S.; Adams, P. D.; Sauter, N. K.; Bergmann, U.; Zouni, A.; Messinger, J.; Kern, J.; Yachandra, V. K.; Yano, J. Untangling the sequence of events during the S2 → S3 transition in photosystem II and implications for the water oxidation mechanism. Proc. Natl. Acad. Sci. U. S. A. 2020, 117, 1262412635,  DOI: 10.1073/pnas.2000529117
    48. 48
      Sanchez-Weatherby, J.; Moraes, I. In The Next Generation in Membrane Protein Structure Determination; Moraes, I., Ed.; Springer International Publishing: Cham, 2016, p 7389.  DOI: 10.1007/978-3-319-35072-1_6 .
    49. 49
      Hellmich, J.; Bommer, M.; Burkhardt, A.; Ibrahim, M.; Kern, J.; Meents, A.; Müh, F.; Dobbek, H.; Zouni, A. Native-like Photosystem II Superstructure at 2.44 Å Resolution through Detergent Extraction from the Protein Crystal. Structure 2014, 22, 16071615,  DOI: 10.1016/j.str.2014.09.007
    50. 50
      Kuo, A.; Bowler, M. W.; Zimmer, J.; Antcliff, J. F.; Doyle, D. A. Increasing the diffraction limit and internal order of a membrane protein crystal by dehydration. J. Struct. Biol. 2003, 141, 97102,  DOI: 10.1016/S1047-8477(02)00633-0
    51. 51
      Russo Krauss, I.; Sica, F.; Mattia, C. A.; Merlino, A. Increasing the X-ray Diffraction Power of Protein Crystals by Dehydration: The Case of Bovine Serum Albumin and a Survey of Literature Data. Int. J. Mol. Sci. 2012, 13, 37823800,  DOI: 10.3390/ijms13033782
    52. 52
      Kwan, T. O. C.; Axford, D.; Moraes, I. Membrane protein crystallography in the era of modern structural biology. Biochem. Soc. Trans. 2020, 48, 25052524,  DOI: 10.1042/BST20200066
    53. 53
      Müh, F.; Zouni, A. Structural basis of light-harvesting in the photosystem II core complex. Protein Sci. 2020, 29, 10901119,  DOI: 10.1002/pro.3841
    54. 54
      Birch, J.; Axford, D.; Foadi, J.; Meyer, A.; Eckhardt, A.; Thielmann, Y.; Moraes, I. The fine art of integral membrane protein crystallisation. Methods 2018, 147, 150162,  DOI: 10.1016/j.ymeth.2018.05.014
    55. 55
      Park, H.; Tran, T.; Lee, J. H.; Park, H.; Disney, M. D. Controlled dehydration improves the diffraction quality of two RNA crystals. BMC Struct. Biol. 2016, 16, 19,  DOI: 10.1186/s12900-016-0069-1
    56. 56
      Francia, F.; Palazzo, G.; Mallardi, A.; Cordone, L.; Venturoli, G. Residual water modulates QA-to-QB electron transfer in bacterial reaction centers embedded in trehalose amorphous matrices. Biophys. J. 2003, 85, 27602775,  DOI: 10.1016/S0006-3495(03)74698-0
    57. 57
      Palazzo, G.; Francia, F.; Mallardi, A.; Giustini, M.; Lopez, F.; Venturoli, G. Water activity regulates the QA to QB electron transfer in photosynthetic reaction centers from Rhodobacter sphaeroides. J. Am. Chem. Soc. 2008, 130, 93539363,  DOI: 10.1021/ja801963a
    58. 58
      Zabelin, A. A.; Khristin, A. M.; Shkuropatova, V. A.; Khatypov, R. A.; Shkuropatov, A. Y. Primary electron transfer in Rhodobacter sphaeroides R-26 reaction centers under dehydration conditions. Biochim. Biophys. Acta, Bioenerg. 2020, 1861, 148238  DOI: 10.1016/j.bbabio.2020.148238
    59. 59
      Noguchi, T.; Sugiura, M. Flash-Induced FTIR Difference Spectra of the Water Oxidizing Complex in Moderately Hydrated Photosystem II Core Films: Effect of Hydration Extent on S-State Transitions. Biochemistry 2002, 41, 23222330,  DOI: 10.1021/bi011954k
    60. 60
      Kaminskaya, O.; Renger, G.; Shuvalov, V. A. Effect of Dehydration on Light-Induced Reactions in Photosystem II: Photoreactions of Cytochrome b559. Biochemistry 2003, 42, 81198132,  DOI: 10.1021/bi020606v
    61. 61
      Pieper, J.; Hauss, T.; Buchsteiner, A.; Baczyński, K.; Adamiak, K.; Lechner, R. E.; Renger, G. Temperature- and Hydration-Dependent Protein Dynamics in Photosystem II of Green Plants Studied by Quasielastic Neutron Scattering. Biochemistry 2007, 46, 1139811409,  DOI: 10.1021/bi700179s
    62. 62
      Beglov, D.; Roux, B. An Integral Equation To Describe the Solvation of Polar Molecules in Liquid Water. J. Phys. Chem. B 1997, 101, 78217826,  DOI: 10.1021/jp971083h
    63. 63
      Kovalenko, A.; Hirata, F. Potential of mean force between two molecular ions in a polar molecular solvent: A study by the three-dimensional reference interaction site model. J. Phys. Chem. B 1999, 103, 79427957,  DOI: 10.1021/jp991300+
    64. 64
      Ben-Shalom, I. Y.; Lin, C.; Kurtzman, T.; Walker, R. C.; Gilson, M. K. Simulating Water Exchange to Buried Binding Sites. J. Chem. Theory Comput. 2019, 15, 26842691,  DOI: 10.1021/acs.jctc.8b01284
    65. 65
      Sindhikara, D. J.; Yoshida, N.; Hirata, F. Placevent: An algorithm for prediction of explicit solvent atom distribution─Application to HIV-1 protease and F-ATP synthase. J. Comput. Chem. 2012, 33, 15361543,  DOI: 10.1002/jcc.22984