Detergent

Article pubs.acs.org/JPCB

All-Atom Molecular Dynamics Simulation of a Photosystem I/Detergent Complex Bradley J. Harris,† Xiaolin Cheng,‡,⊥ and Paul Frymier*,†,§,∥ †

Department of Chemical and Biomolecular Engineering, ‡Department of Biochemistry and Cellular and Molecular Biology, Sustainable Energy Education and Research Center, and ∥Bredesen Center for Interdisciplinary Research and Graduate Education, University of Tennessee, Knoxville, Tennessee 37996, United States ⊥ Center for Molecular Biophysics, Oak Ridge National Laboratory, Oak Ridge, Tennessee 37831, United States §

S Supporting Information *

ABSTRACT: All-atom molecular dynamics (MD) simulation was used to investigate the solution structure and dynamics of the photosynthetic pigment−protein complex photosystem I (PSI) from Thermosynechococcus elongatus embedded in a toroidal belt of n-dodecyl-β-D-maltoside (DDM) detergent. Evaluation of root-mean-square deviations (RMSDs) relative to the known crystal structure show that the protein complex surrounded by DDM molecules is stable during the 200 ns simulation time, and root-mean-square fluctuation (RMSF) analysis indicates that regions of high local mobility correspond to solvent-exposed regions such as turns in the transmembrane α-helices and flexible loops on the stromal and lumenal faces. Comparing the protein−detergent complex to a pure detergent micelle, the detergent surrounding the PSI trimer is found to be less densely packed but with more ordered detergent tails, contrary to what is seen in most lipid bilayer models. We also investigated any functional implications for the observed conformational dynamics and protein−detergent interactions, discovering interesting structural changes in the psaL subunits associated with maintaining the trimeric structure of the protein. Importantly, we find that the docking of soluble electron mediators such as cytochrome c6 and ferredoxin to PSI is not significantly impacted by the solubilization of PSI in detergent.

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INTRODUCTION Membrane proteins comprise approximately 30% of open reading frames, highlighting their biological importance.1 However, they represent only a small, although rapidly increasing, fraction of known protein structures, reflecting the difficulties of structural determination of membrane proteins.2 This problem is partially due to the need for solubilization of the transmembrane surface via a detergent or lipid assembly. These detergents disrupt the native cell membrane and solubilize the membrane proteins in mixed micelles that prevent aggregation by mimicking the native membrane environment.3,4 Unfortunately, little is known of the aqueous structure and protein−detergent interactions of solubilized membrane proteins, and how these detergents may interfere with the structural and functional properties of these proteins.5,6 Photosystem I (PSI) is a large protein complex involved in cyanobacterial, algal, and plant photosynthesis. It provides both large antennae for harvesting solar energy and a reaction center for accomplishing stable charge separation. In the cyanobacterium Thermosynechococcus elongatus (T. elongatus), PSI is located in the thylakoid membrane and exists primarily as a homotrimer of three 12-subunit monomers, each of which contains several membrane-spanning domains. The crystal structure of trimeric PSI from the thermophilic cyanobacterium T. elongatus has been resolved at 2.5 Å7 (Figure 1) as a large complex (∼1 MDa) containing the reaction center pigments, © 2014 American Chemical Society

light harvesting chlorophyll a (chl a), carotenoids, quinones, and iron−sulfur clusters that mediate electron transport to support carbon fixation and other redox reactions.8 The ability of PSI to capture photons as well as its stable nature has led to many studies investigating in vitro applications for alternative energy solutions, such as hydrogen or direct electricity production.9−18 For efficient energy conversion, it is important to develop robust, reproducible techniques to control the orientation of PSI molecules on conductive substrates or in complex with other proteins, which requires better knowledge of detergent−protein interactions in order to avoid denaturation and/or random orientation of PSI.19 This knowledge will also prove important in the engineering of genetic modifications, such as binding motifs necessary for highly specific binding of PSI to surfaces or partner proteins, in order to ensure the binding is not hindered by the presence of detergent. Overall, a better understanding of PSI−detergent interactions could help develop technologies that prolong the lifetime of these biobased energy conversion devices by reducing unbound or inactive proteins. Molecular dynamics (MD) simulation is a useful computational technique for the study of biomacromolecules.20−23 It has been employed to study the conformational dynamics of Received: July 17, 2014 Revised: September 18, 2014 Published: September 18, 2014 11633

dx.doi.org/10.1021/jp507157e | J. Phys. Chem. B 2014, 118, 11633−11645

The Journal of Physical Chemistry B

Article

Figure 1. (A) Transmembrane view of the crystal structure of monomeric PSI from T. elongatus (PDB ID: 1JB0) with the stromal side above and lumenal side below. (B) Top view looking down on the lumenal face. In both images, the protein is shown in ribbons format with the reaction center subunits (psaAB) in green, the terminal electron acceptor (psaC) in red, the exterior stromal subunits (psaDE) in blue, and the peripheral transmembrane helices (psaFIJKLMX) in orange.

pure lipid bilayers,24−26 as well as peptides and proteins embedded in lipid bilayers, such as the porin OmpF,27,28 the outer membrane protein OmpA, 29 the water channel aquaporin,30,31 the ion channels gramicidin A,32,33 KscA,34,35 and MscL,36,37 G-protein-coupled receptors (GCPRs),38−40 as well as purple bacterial chromatophores41−43 and the photosynthetic protein−cofactor complex photosystem II (PSII),44 among others. MD simulation has also been conducted on a variety of detergent micelles,45−52 and on membrane protein− detergent systems. In 2004, researchers demonstrated spontaneous membrane protein−detergent micelle formation for both a simple α-helical membrane protein glycophorin A (GpA)53 and the outer membrane protein OmpA54 using MD simulation. More recently, MD was used to study the structure and dynamics of the channel protein FhuA embedded in a detergent belt,55 and to build a model of aquaporin-0 surrounded by detergent molecules for comparison to smallangle X-ray scattering data.56 Recently, we investigated the structure of PSI trimer solubilized with n-dodecyl-β-D-maltoside (DDM) detergent using small-angle neutron scattering (SANS) combined with MD simulations.57 DDM is a mild, nondenaturing detergent, commonly used in protein extraction and purification, which was used for the crystallization of this complex.7 Here we report on all-atom MD simulations conducted on photosystem I (PSI) trimer from T. elongatus surrounded by a monolayer toroidal belt of DDM. We have performed extensive simulations for indepth analysis of the structure and dynamics of the PSI trimer embedded in a detergent environment. With these simulations, we are able to examine the following issues to a level of detail not previously possible: stability and flexibility of the PSI trimer in a detergent environment, detailed protein−detergent interactions and their mutual effects on each other’s behavior, and possible implications of the observed conformational dynamics for the function of this photoactive pigment−protein complex.

hydrophilic head groups pointed outward. A base case of 132 DDM molecules was chosen on the basis of the reported aggregation number for this detergent,49 with micelles of 70 and 200 DDM molecules also simulated as extreme cases. All systems were subjected to 20 ps of MD equilibration in vacuo, and subsequently solvated with TIP3P water molecules. These equilibration steps were conducted to remove clashes between atoms, heat the system gradually to the target temperature, and reach a kinetic energy equi-partition in the system. This is meant to ensure that the systems do not explore unrealistically high-energy conformational spaces prior to MD production runs. Solvated systems were subjected to an additional 20 ps of MD equilibration, and then 100 ns MD production runs were carried out. Setup of PSI−DDM Micelle Systems. The 2.5 Å PSI crystal structure from T. elongatus (PDB ID: 1JB0) was chosen as the starting model.7 The PSI trimer was generated using the transformation matrix contained therein. There are 91 out of a total of ∼2300 residues missing from the crystal structure of monomeric PSI which were not included in the simulations. In general, these unresolved regions span less than 15 successive residues, except in the cases of PsaF and PsaK, which are missing 23 and 54 amino acids from their N-termini, respectively. The missing residues are terminally located and are likely flexible. These intrinsically disordered regions are expected to play a role in PSI−mediator interactions but are unlikely to significantly affect the global dynamics of the PSI/ DDM complex during the simulation. Phylloquinone molecules located in the reaction center core (two per PSI monomer) were not included in the simulation. A monolayer belt of DDM detergent was built around the periphery of PSI consisting of semicircular planes of DDM densely packed around the hydrophobic exterior transmembrane surface of the protein. Additional DDM molecules were placed in the interstitial voids between individual PSI monomers in a bilayer orientation, resulting in the so-called void-filled ring model, the analysis of which is the focus of this paper. An alternative random model was generated by randomly placing DDM molecules around the PSI trimer using Packmol,58 and removing all those which overlapped with the protein complex. For the model including the associated

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METHODS Setup of Pure DDM Micelle Systems. Pure DDM micelles were built using Packmol,58 constrained to a spherical orientation with the hydrophobic tails pointed inward and the 11634



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lipids 1,2-distearoyl-monogalactosyldiglyceride (LMG) and 1,2dipalmitoyl-phosphatidylglycerol (LHG), the structure and coordinates of all identified lipids were taken from the crystal structure and atom types and parameters from the updated CHARMM force field for lipids.59 All systems were subjected to 50 ps of MD equilibration in vacuo, and subsequently solvated with TIP3P water molecules. Sodium and chloride ions were then added by random replacement of water molecules in order to neutralize each system. Solvated systems were subjected to an additional 50 ps of MD equilibration, and then, MD production runs were carried out. Simulation Details. All simulations were carried out using NAMD,20 with an extended version of the CHARMM27 force field.60 Parameters for chlorophyll a and beta-carotene were taken from parametrization of photosystem II (PSII) cofactors.61 Parameters for the iron−sulfur clusters were based on the work done by Smith and co-workers,62 and those for DDM were developed by Abel et al.49 All simulations were performed at constant temperature (310 K), pressure (1 atm), and number of particles. Electrostatics were calculated using the particle mesh Ewald63 method with a 1 nm cutoff for the real space calculation. A cutoff of 1 nm was also used for van der Waals interactions. System temperature was maintained by controlling the kinetic energy of the system using Langevin dynamics, with a damping coefficient of 10 ps−1. System pressure was controlled using the Langevin piston method,64 with an oscillation time constant of 200 fs and a damping time constant of 100 fs. The time step for integration was 2 fs for all simulations, and coordinates and velocities were saved every 20 ps. Data analyses used either VMD65 or locally written code. All images were rendered in VMD. Predicted Docking of Soluble Electron Mediators. Docking predictions for cytochrome c6 (PDB ID: 1C6S) and ferredoxin (PDB ID: 2CJN) to PSI were conducted on the ZDOCK online server.66 Model predictions were generated for every monomer of trimeric PSI using snapshots of the MD trajectory taken for every 20 ns of simulation; predictions based on the PSI crystal structure were also generated as a reference. For cytochrome c6, only the psaAB subunits of PSI were uploaded to the server, and W655A and W631B were specified as contact residues.67 In the case of ferredoxin, only the psaCDE subunits of PSI were considered, and I11C, T14C, Q15C, K34C, K104D, and R39E were specified as contact residues.68 Cα RMSD values of the top 10 predicted bound mediator structures (if available) were calculated for each time step relative to the crystal structure predictions for each of the three monomers of the PSI trimer, with the minimum Cα RMSD for each time step being reported.

Figure 2. Initial configuration of the protein−detergent simulation system: (A) top view looking down at the lumenal face; (B) transmembrane view with the stromal side above and lumenal side below. The detergent molecules are shown in lines format, with the carbon atoms in cyan and the oxygen atoms in red. Water molecules and counterions are omitted for clarity.

with the hydrophobic tails pointed inward, in order to fill the interstitial hydrophobic voids between individual PSI monomers. The aggregation number for this system has been previously estimated to be ∼790 DDM molecules per PSI trimer, assuming uniform packing of detergent around the transmembrane region.57 Because this aggregation number does not account for the void space between individual PSI monomers, we chose to use 1000 DDM molecules in our simulations, with 800 detergent monomers in a toroidal belt around the transmembrane domain and 200 in the interstitial voids. Subsequently, this system was energy-minimized, solvated, and equilibrated (see the Methods section). The equilibrated system was followed by a 200 ns production run. A theoretical scattering curve generated on the basis of this protein−detergent complex after 50 ns of MD simulation was found to reproduce experimental SANS data with a χ2 value of 3.83.57 Protein Dynamics. Protein Stability. In order to analyze the overall stability of the PSI−DDM complex, we calculated the root-mean-squared deviation (RMSD) of the PSI trimer relative to the initial crystal structure throughout the course of the simulation (Figure 3). The Cα RMSD calculated for all residues rose to ∼2.5 Å after ∼50 ns, drifting to ∼2.6 Å after ∼100 ns and remaining there for the duration of the simulation. This extended drift phenomenon has also been observed in a previous MD study of OmpA in dodecylphosphocholine (DPC) detergent.70 A comparable backbone RMSD value of ∼2.0 Å has previously been reported in a 50 ns MD simulation of FhuA in N-octyl-2-hydroxyethyl sulfoxide (OES) detergent.55 These values fall within the same range as MD studies of membrane proteins in lipid bilayers. For example, 15−20 ns of MD simulations of OmpA in a dimyristoyl phosphatidylcholine (DMPC) bilayer and KcsA in a palmitoyl oleoyl phosphati-

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RESULTS AND DISCUSSION The PSI−DDM complex is known to be disk-like, 30 nm in diameter by 9 nm in height, as determined from dynamic light scattering.69 This experimental evidence, combined with the fact that a simulation of randomly placed DDM molecules around the PSI trimer had not begun to converge after >20 ns of MD (data not shown), led us to use a preformed disk-like protein−detergent assembly as our starting structure. The initial PSI−detergent system is depicted in Figure 2. We used the characteristic hydrophobic belts of membrane proteins as a guide to constructing a uniform, toroidal belt of DDM detergent around the transmembrane domain of the PSI trimer. We added additional DDM molecules into the interstitial space of the PSI trimer, in a bilayer orientation 11635



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Figure 3. Cα RMSD values versus time for the PSI−DDM micelle simulation. All curves were generated from the simulation starting structure. The lines show the Cα RMSD values for: all residues (black continuous line); the reaction center subunits psaA and psaB (gray continuous line); the terminal electron acceptor psaC (black dotted line); the ferredoxin docking subunits psaD and psaE (black dashed line); and the peripheral transmembrane helices psaF, psaI, psaJ, psaK, psaL, psaM, and psaX (gray dotted line).

dylcholine (POPC) bilayer resulted in Cα values of 2.0 and 2.3 Å, respectively.71 Similar values were found for a 10 ns simulation of PSII embedded in the thylakoid membrane.44 These results suggest that the protein in the micellar environment shows a similar magnitude of structural drift from the crystal structure as that in a lipid bilayer. We verified that a stable conformation had been reached by calculating the Cα RMSD of the PSI trimer versus a snapshot of the complex after 100 ns of MD simulation, which reached a plateau of 1.5 Å (Figure S1, Supporting Information). We further analyzed the protein structural drift by decomposing the RMSD values into those of various individual structural components of PSI (highlighted in Figure 1). For this purpose, we divided the structure into four domains: subunits psaA and psaB, which house the light-harvesting reaction center and associated pigments; subunit psaC, found on the stromal face and housing the terminal electron acceptors FA and FB;7 subunits psaD and psaE, also located on the stromal face and believed to be involved in the docking of ferredoxin;72 and the peripheral transmembrane subunits psaI, psaJ, psaK, psaL, psaM, psaX, as well as psaF, which traverses the membrane and is believed to facilitate docking of both ferredoxin on the stromal face as well as of cytochrome c6 on the lumenal face.8 We found that the reaction center subunits psaA and psaB display the lowest structural drift with a final Cα RMSD value of ∼1.8 Å. This can be explained by the fact that these subunits represent the core of the protein and have little exposure to the solvent, which is consistent with the root-mean-square fluctuation (RMSF) analysis discussed below. The terminal electron acceptor subunit psaC exhibits Cα RMSD values in the range 1.5−2.5 Å, albeit with greater fluctuations. This subunit is located on the stromal surface, and therefore has little interaction with detergent but is highly exposed to the solvent. In contrast, the peripheral transmembrane helices exhibit the greatest drift with a maximum Cα RMSD value of ∼4.0 Å. This plateau is reached after 125 ns of MD simulation and is stable for the remainder of the 200 ns simulation time, indicating this region has reached equilibrium. This substantial deviation is likely due to interactions with surrounding detergent molecules in this system, an environment that is quite different from the low temperatures and tight helical packing necessary for obtaining a high-resolution crystal structure. As is the case for psaK (shown in Figure 4A and B), these subunits bend and

Figure 4. Simulation snapshots of the peripheral transmembrane αhelix psaK (A) prior to MD simulation; (B) after 200 ns of MD simulation. PsaK is shown in ribbons format in blue with detergent molecules within 5 Å of psaK shown in surface format with the head groups in red and the tail groups in cyan. Snapshots of the core subunits psaA and psaB of PSI monomer and the surrounding detergent belt (C) prior to MD simulation; (D) after 200 ns of MD simulation. The psaAB subunits are shown as blue ribbons, the psaCDE subunits as red ribbons, and the DDM molecules are shown in licorice format with the heads in red and the tails in cyan.

contort during the simulation, becoming more compact and kinked as the detergent molecules surrounding them evolve from the initial uniform belt. Chandler et al. observed similar results in their 20 ns MD study of a model chromatophore complex of light harvesting complex LH1 and the reaction center (RC) embedded in a lipid bilayer, wherein they reported RMSD values of ∼4 Å for the core RC protein and ∼8 Å for the surrounding LH1 domain.41 Additionally, Cα RMSD values in the range 5−9 Å have previously been reported in MD studies of the membrane proteins Mistic73 and aquaporin56 in detergent micelles. The outer stromal subunits psaD and psaE show unique behavior, with the Cα RMSD rising quickly to ∼2.5 Å after ∼15 ns, before climbing to ∼3.0 Å after ∼50 ns and fluctuating between 2.5 and 3.5 Å for the rest of the simulation. This may be explained by the shrinking of the detergent belt over the course of the simulation, as it conforms to the nonuniform hydrophobic periphery of the PSI trimer, moving detergent molecules farther away from the stromal domains (Figure 4C and D). It should be noted that individual detergent molecules exhibited significant translational mobility in both the protein−detergent micelle and pure micelle simulations but that no loss of detergent from the complexes was observed. Protein Flexibility. We also calculated the time-averaged root-mean-square fluctuation (RMSF) for each residue (Figure 5). RMSF is a measure of the deviation of backbone atoms (N, Cα, and C atoms) from their average positions during the MD simulations, providing insight into the thermal fluctuations and 11636



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Figure 5. (A) Broken-line plot of time-averaged Cα RMSF values versus residue number for PSI (solid black line), calculated over the last 100 ns of the PSI/DDM MD simulation, compared with RMSF values calculated from the temperature factors of the X-ray structure (dotted black line). RMSF values were calculated for each individual monomer of the PSI trimer and then averaged, with error bars representing the standard deviation. Residues corresponding to the reaction center (RX CTR), stromal (S), and peripheral transmembrane (PT) domains are labeled correspondingly. (B) A zoomed-in view of time-averaged Cα RMSF values versus residue for the stromal domain.

Figure 6. (A) Top view of the entire PSI trimer, shown in blue ribbons. The psaL subunits are shown in red, with the N-termini highlighted in boxes. (B) Transmembrane view of the psaA (blue) and psaL (red) subunits after 40 ns of MD simulation with the associated lipids identified in the crystal structure included, shown in ribbons format with the associated phospholipid shown in VDW format. (C) Transmembrane view of the psaA and psaL subunits after 200 ns MD simulation without the associated lipids identified in the crystal structure.

experimentally, as can be seen in both the reaction center and peripheral transmembrane domains. This is also the case for solvent-exposed loops located on the stromal face (Figure 5B). Most noticeable is the extremely high RMSF value of ∼7.4 Å obtained for the N-terminus of psaL. This is due to the extension of this flexible region from the confines of the interstitial void out into the solvent over the course of the simulation, which may be due to the absence of the associated lipids from the simulation and will be discussed in detail below; this occurrence is shown in detail in Figure 6. Similarly high RMSF values for the N- and C-termini of transmembrane helices were observed in MD of aquaporin-0 in a DMPC

atomic mobility of proteins and protein complexes. For comparison, we also converted the B-factors of the 2.5 Å resolution crystal structure to equivalent RMSF values. The overall trends of the experimental and simulated curves are qualitatively similar, and there appears to be a correlation between local mobility and protein structure. Regions of low local mobility are primarily confined to the core residues of the reaction center subunits which are not solvent-exposed. The calculated RMSF values are virtually identical in these regions to those obtained from experiment. In contrast, the RMSF values for turns in the transmembrane α-helices obtained from the simulation are significantly higher than those determined 11637



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Table 1. Simulation Details system PSI−DDM void-filled ring model PSI−DDM w/ lipids model PSI−DDM random model DDM micelle

no. of water molecules

no. of detergents

PSI, DDM

248 861

1002

PSI, DDM, LMG, LHG PSI, DDM

245 775

1002

912 442

997

26 116

132

components

DDM

no. of lipids

3 LHG, 1 LMG

no. of ions

no. of atoms

box size (nm)

simulation time (ns)

44 Na+

977 756

28.0 × 28.0 × 13.5

200

283 Na+, 230 Cl− 44 Na+

970 497

28.0 × 28.0 × 13.5

40

2 986 094

35.0 × 35.0 × 25.0

20

89 040

10.0 × 10.0 × 10.0

100

Figure 7. (A) Side view of the PSI/DDM micelle prior to MD simulation and (B) after 200 ns of MD simulation. (C) Side view of the pure DDM micelle prior to MD simulation and (D) after 100 ns of MD simulation. In all cases, the detergent is shown in blue in low resolution surface representation, with the protein shown in red ribbons; water and counterions are omitted for clarity. Note: these images are not to scale.

bilayer.74 Overall, regions confined to the interior of the protein or shielded by detergent exhibit low local mobility, with higher RMSF values seen in loops or turns exposed to the solvent. Micelle Dynamics. Micelle Shape. Having examined the stability and mobility of the PSI−DDM complex, we were further interested in comparing the behavior of detergent in this system to a pure DDM micelle in order to elucidate the effect of the protein on detergent behavior. To accomplish this, a 100 ns MD simulation was conducted for a 132-detergent DDM micelle, the previously reported aggregation number for pure DDM in water,49 for comparison (simulation details for each case are shown in Table 1). To begin, we visually inspected changes in the micelle shape for each system throughout the course of MD. Snapshots of the PSI−DDM micelle simulation are shown in Figure 7A and B below. The thickness of the detergent belt shrinks during dynamics, conforming to the nonuniform hydrophobic periphery of the protein. Overall, the protein−detergent micelle becomes more ellipsoidal in shape over the course of the simulation. The pure detergent micelle was initially constructed in a spherical configuration but also transitions to an ellipsoid over the course of MD simulation (Figure 7C and D). Interfacial Properties. The packing of detergent molecules, and the resulting extent of water penetration, is an additional interesting aspect of these systems, as it can influence the

internal dynamics of micellar aggregates as well as the protein assembly embedded in the micelle. Radial density profiles (relative to the center of mass) of the two systems (Figure 8) show that the detergent atom distributions and the solvent− detergent interface are quite broad, as observed in previous studies. Protein−detergent interactions in the PSI−DDM complex result in a shift in the distributions of the detergent head and tail atoms compared to the pure detergent micelle. In the pure micelle system, the atomic density profile of the detergent tail is asymmetric, peaking sharply at 1.0 Å before gradually reducing toward the center of the micelle, with a width of ∼30 Å between each minimum; the remaining density at the center of the aggregate illustrates the tightly packed nature of the detergent tails. In the PSI−DDM system, the tail atom density becomes more symmetric with a peak at 100 Å that reduces to zero by 80 Å due to the presence of protein at the center of the complex, and also broadens to a width of ∼40 Å. The head atom density distribution exhibits the reverse trend, with a symmetric profile peaking at 29 Å in the case of the DDM micelle, and an asymmetric curve for the PSI−DDM complex with a peak at 119 Å; the width of the distribution also broadens from ∼35 Å for the DDM micelle to ∼50 Å in the case of PSI−DDM. In both systems, the lipid tails are more localized, exhibiting high peak density values, whereas the headgroup peak densities are less pronounced. A similar trend 11638



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systems. Similar results were seen in MD simulations of DDM micelles in water.49 For the PSI−DDM system, the nonzero water atom density throughout the protein is due to the penetration of water into cavities on the stromal and lumenal faces. Another interesting comparison between these two systems comes from analyzing the distance between the head and tail density peaks. This value decreases from 28 Å for the pure micelle system to 19 Å for the protein−detergent micelle, indicating that the head-to-tail length of the detergent molecules is shrinking in order to match the detergent belt thickness to that of the hydrophobic transmembrane region of the PSI trimer. This process can clearly be seen in Figure 4, where the DDM molecules surrounding subunits psaA and psaB depart from the initial belt configuration as they shrink and conform to the transmembrane region of the protein over the course of the simulation. We also calculated the solvent-accessible surface area (SASA) of the detergent atoms for both the protein−detergent complex and the pure detergent micelle (Table 2). The calculation of SASA involves drawing a mesh of points by extending the known radius of each atom of a given molecule by the radius of a water molecule; these points are then checked against the surface of neighboring atoms to determine if they are buried or accessible. The number of points accessible is then multiplied by the portion of surface area each point represents to determine the SASA; it can be used to evaluate water accessibility and thus detergent packing. The higher SASA value of 285.7 ± 9.6 Å2 calculated for the PSI−DDM complex, compared to 247.6 ± 12.3 Å2 for a pure DDM micelle, indicates a slightly more diffuse arrangement of detergent molecules in the former. We further calculated the SASA for four distinct groupsthe first glucose ring of the detergent headgroup, the second glucose ring of the headgroup, the first six atoms of the hydrocarbon tail, and the last six atoms of the tail (a single DDM molecule is shown with Table 3 as a reference)and found that the SASA values for the head atoms are actually lower for the protein−detergent complex compared to the pure detergent micelle, while the tail atom values are higher. This shows that the head groups are more tightly packed in the PSI− DDM system compared to the pure DDM micelle, while the tail groups are more loosely arranged. These results would suggest that the detergent belt surrounding PSI is becoming more like a bilayer structure than a micelle structure over the course of the simulation. Plots versus time for both the PSI/ DDM complex and the pure DDM micelle show that the SASA quickly reaches an equilibrium value, and further confirm the stability of these systems during MD simulation (Figure S3, Supporting Information). Internal Structure and Dynamics. The dihedral order parameter S2 is a measure of the equilibrium distribution of the orientation of hydrocarbon chains, and provides information on the chain conformations and fluctuations in the micelle

Figure 8. Radial atom density distributions for the various system components of (A) the PSI−DDM complex and (B) the pure DDM micelle. In each case, the atom densities of the components (protein = solid black line; hydrophobic tails = dotted black line; hydrophilic heads = dotted gray line; water = solid gray line) are plotted as a function of the distance from the center of mass of the system. Note that the water density is plotted on a separate scale in part B.

was observed for MD simulations of aquaporin-0 in a DMPC bilayer.74 We have also plotted the density curves of the detergent head and tail atoms before and after MD simulation for both the pure DDM micelle and PSI/DDM complex (Figure S2, Supporting Information). In the case of the pure micelle, the tail atomic density curve sharpens over the course of MD as the detergent tails shift from the initial configuration to become more tightly packed, while the head atomic density peak moves outward, possibly due to the transition from a spherical to an ellipsoidal micelle. A similar sharpening of the tail atomic density curve is observed over the course of MD in the PSI/DDM complex, while changes in the head atomic density profile are minimal. Examining the atomic density profiles at various time points, we also find that the radial distances corresponding to the maximum head and tail atomic densities are stable throughout the course of the simulation for both the PSI−DDM and pure DDM micelle systems (Table S1, Supporting Information). The water atom density is similar in both systems, approaching the bulk density value before the detergent head atom density curve has reached zero, indicating solvation of the hydrophilic micelle headgroups. In other words, water significantly penetrates the DDM detergent in both the protein−detergent complex and pure detergent micelle Table 2. Solvent-Accessible Surface Area per Detergenta system

total (Å2)

1st headb (Å2)

2nd headc (Å2)

upper taild (Å2)

lower taile (Å2)

PSI−DDM complex DDM micelle

285.7 ± 9.6 247.6 ± 12.3

213.7 ± 1.3 224.0 ± 3.1

195.9 ± 1.5 201.7 ± 3.7

154.1 ± 1.7 148.2 ± 4.0

125.9 ± 1.7 98.1 ± 3.3

a For each system, SAS was calculated using the VMD plugin with a probe radius of 1.4 Å. bThe first six carbon and five oxygen atoms (first glucose ring). cThe second six carbon and oxygen atoms (second glucose ring). dThe first six carbon atoms of the hydrocarbon tail. eThe last six carbon atoms of the hydrocarbon tail.

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Table 3. Comparison of the Properties of the Detergent Taila

dihedral angle order parameter (S2) carbon no. 1 2 3 4 5 6 7 8 9 10

b

PSI−DDM 0.057 0.049 0.050 0.052 0.047 0.045 0.042 0.037 0.027 0.020

± ± ± ± ± ± ± ± ± ±

0.012 0.014 0.014 0.012 0.013 0.013 0.013 0.012 0.012 0.011

MSD of tail hydrogen atoms (Å2)

DDM 0.026 0.020 0.019 0.025 0.021 0.021 0.018 0.015 0.016 0.006

± ± ± ± ± ± ± ± ± ±

PSI−DDM

0.036 0.033 0.035 0.033 0.033 0.031 0.037 0.032 0.030 0.032

58.0 56.6 56.0 55.6 55.5 55.8 56.4 57.6 59.2 61.5

± ± ± ± ± ± ± ± ± ±

22.5 21.9 21.6 21.4 21.2 21.2 21.3 21.7 22.4 23.7

DDM 84.7 86.0 85.5 86.1 86.6 88.0 89.4 91.7 94.3 97.7

± ± ± ± ± ± ± ± ± ±

33.4 33.8 33.6 33.8 33.8 34.3 34.7 35.7 36.8 38.7

a

These values were time-averaged for the last 100 ns (out of 200 ns) and the last 50 ns (out of 100 ns) for the PSI/DDM and pure DDM systems, respectively. bNumbered from the first (closest to the headgroup) to last CH2 group of the detergent tail.

interior.75 Values for this parameter vary from 0.0 to 1.0, covering the spectrum from random, uninhibited fluctuations to rigidly fixed conformations. Time-averaged S2 values for the tail CH2 groups are consistently higher for the PSI−DDM complex compared to pure DDM (Table 3), indicating that the presence of protein results in more ordered detergent tail structures. This agrees with the favorable interactions we have noted between the detergent and the hydrophobic periphery of PSI, both through visual inspection of the thinning of the detergent belt as it conforms to the protein and through quantification of the shrinking of the average length of the DDM molecule compared to a pure DDM micelle. Also, noticeable is the fact that the variance (standard deviation) over time for the pure DDM micelle is roughly equal to the mean, reflecting the greater fluctuation of this system; this value is smaller in the case of the protein−detergent complex, with the standard deviation equating to ∼20% of the mean. We have also analyzed diffusion of the detergent tail by calculating the timeaveraged mean-square displacements (MSDs) for the hydrogen atoms of each tail CH2 group. We again conclude that the detergent tails are less diffusive (more ordered) in the PSI− DDM system, as the MSD values are lower for the PSI−DDM complex compared to the pure DDM micelle (Table 3). These results, combined with the SASA calculations, indicate that the detergent is overall more loosely packed in the protein− detergent complex than in a pure detergent micelle but that the detergent tails are more ordered. This is contrary to what is typically seen in most lipid bilayer models, where the liquid

disordered phase is more loosely packed than the liquid ordered phase.76,77 Implications for Function. As referenced previously, PSI is an integral part of the photosynthetic cycle of plants and microorganisms, harnessing solar energy in order to accomplish electron transfer across the thylakoid membrane. Having shown that our MD model of this protein embedded in a detergent ring is a stable complex, we were further interested in discerning any potential effects of solution dynamics and protein−detergent interactions on the superstructure and function of PSI. Changes in Trimeric Structure of PSI. PSI is known to exist as a trimer in cyanobacteria, while adopting a monomeric structure in algae and higher plants. In plant PSI, the peripheral interfaces are involved in interactions with additional peripheral antenna, including the light-harvesting complex II (LHCII).78 Functionally, LHCII recruitment is vital for regulation and protection from high light intensity and photodamage common to the land surface and shallow water environments that these organisms are typically found in. In comparison, cyanobacterial PSI does not possess an external antenna system and the peripheral interfaces of the protein are involved primarily in trimer-maintaining interactions. The function of the trimer could be to provide a larger antenna system for optimal capture of dim light, as low light conditions are common to the natural habitat of these species.79 PsaL and its associated chlorophylls are believed to be central to this process, serving as sites for excitation energy transfer between adjacent monomers in a 11640



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trimer. Indeed, growth of T. elongatus psaL deletion mutants decreased by a factor of 10 under low light compared to wild type, and were shown to be unable to form trimers.80 One intriguing result of this simulation study is the high local mobility of the N-terminus of psaL, resulting from its extension from the trimer void out into the solvent (shown in Figure 6C). This extension occurred for two out of the three psaL subunits of the trimer and is a relatively fast event, occurring after ∼20 ns of MD simulation. Crystallized trimeric PSI has been found to be associated with several lipids, with one particular phospholipid being bound to psaA and in close proximity to psaL and the monomer−monomer interface.7 In order to further investigate, we conducted MD simulations of the PSI/ DDM complex with the associated lipids identified in the crystal structure included, and found that the N-terminus of all three psaL subunits remained in all trimer voids without extending for the entirety of a 40 ns simulation (Figure 6B). The absence of this lipid from our MD simulations could contribute to the extension of the psaL N-terminus during dynamics, and could result in further destabilization of the trimeric structure on longer timescales, as psaL is crucial to the trimerization process and these integral lipids are believed to be functionally important in the PSI complex and not mere preparation artifacts. Similarly associated phospholipids have been found to be functionally important in the case of the yeast cytochrome bc(1) complex81 and the photosynthetic reaction center of the purple bacterium Rhodobacter sphaeroides.82 Docking of Soluble Electron Mediators. The function of PSI in the photosynthetic cycle involves the harnessing of solar energy in order to accomplish the transfer of electrons provided by a soluble electron mediator such as cytochrome c6 (cyt c6) on the lumenal side of the thylakoid membrane to ferredoxin (Fd) on the stromal side. These electrons will eventually be used to reduce NADP+ to NADPH, thereby providing reducing equivalents for the cell.8 Researchers have thoroughly investigated the docking of cyt c6 and plastocyanin (PC) on the lumenal face of PSI, as well as of Fd on the stromal face. Sommer et al. identified the major interaction site for cyt c6/PC docking as a hydrophobic lumenal indentation formed by the psaA and psaB subunits and further discovered two tryptophan residues vital to this process,67 while mutagenesis studies conducted by several research groups have identified residues of the psaC, psaD, and psaE stromal ridge subunits that are critical to Fd docking on the stromal face due to electrostatic interactions.68 All-atom MD studies have been previously conducted to observe the docking of cytochrome c6 and plastocyanin to PSI monomer in vitro.83 Using the docking prediction server ZDOCK,66 we have investigated the docking of cyt c6 and Fd to PSI. By comparing docking predictions obtained for snapshots of the MD simulation trajectory to that of the crystal structure, we were able to assess the effects of solution dynamics and protein− detergent interactions on these docking processes. Overall, we found that the presence of detergent had little effect on the docking of soluble electron mediators, as the predicted models for MD snapshots did not differ significantly from those obtained for the crystal structure. This was evaluated visually and by calculating Cα RMSD values of predicted docked mediator structures during dynamics relative to the predicted docked structures using the PSI crystal structure (Table 4). Cyt c6 docking to MD simulation snapshots differed by an average of 2.31 ± 1.01 Å relative to the crystal structure over the course of the simulation, while Fd docking differed by an average of

Table 4. Cα RMSD Values for Mediator Docked to PSI/ DDM MD Simulation Snapshots time (ns)

Cyt c6 docking RMSD (Å)

Fd docking RMSD (Å)

20 40 60 80 100 120 140 160 180 200

1.50 3.58 1.21 2.20 1.55 3.81 3.72 2.11a 1.63 1.77

5.25 1.98 2.60 3.07 3.18 3.07 4.28 3.38 2.83 2.14

a Predicted docking of cyt c6 was successful for only two of three PSI monomers for the 160 ns snapshot.

3.18 ± 0.97 Å. We believe these slight differences in binding can be attributed to fluctuations in key binding site residues, suggesting an induced-fit mechanism for mediator docking. For the case of ferredoxin, the changes in mediator docking can likely be explained by the ∼3.0 Å structural drift of the stromal psaDE subunits relative to the crystal structure during the simulation (Figure 2), resulting in conformational changes that alter the binding of Fd. We also note that several residues believed to be associated with Fd docking exhibit high fluctuations during the simulation, specifically residues T14 and Q15 of psaC and residue K104 of psaD (Table 5). Table 5. RMSFs for PSI−Mediator Binding Site Residues residuea stromal

lumenal

T14C Q15C K104D R627B D628B Y629B L630B

PSI−DDM MD model RMSF (Å)

crystal structure RMSF (Å)

± ± ± ± ± ± ±

1.26 1.25 1.13 1.07 1.13 1.17 1.13

1.96 2.00 2.74 1.61 1.64 1.89 1.60

0.02 0.03 1.13 0.69 0.50 0.56 0.55

a The format for the residue identifier is a one-letter amino acid symbol, residue number, and subunit name.

Mutations of these residues in PSI are known to affect the charge of the binding pocket for Fd.68 Furthermore, the triple mutant I12V/T15K/Q16R of psaC in PSI from the green algae Chlamydomonas reinhardtii, which corresponds to I11V/T14K/ Q15R in T. elongatus, has been shown to reduce the binding of Fd by 2 orders of magnitude.84 Thus, fluctuations in these key residues could be contributing to the altered binding of ferredoxin to PSI in detergent-solubilized conditions. In the case of cytochrome c6, the explanation is not so clear, as this mediator binds to a hydrophobic indentation formed by the psaAB subunits on the lumenal face of PSI and these reaction center subunits exhibit the lowest RMSD values relative to the crystal structure (Figure 2). Furthermore, the residues believed to be involved in binding of cyt c6, W655 from psaA and W631 from psaB, do not fluctuate drastically. However, there are several residues of the lumenal helix l of psaB, namely, R627 through L630, that exhibit unusually high RMSFs (Table 5). Mutagenesis studies have shown that helix l is essential for binding and electron transfer with the soluble electron donors cytochrome c6 and plastocyanin.85 Therefore, 11641



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fluctuations in this luminal helix loop could cause the changes in binding of cyt c6 to PSI that we have observed.

have an impact on the trimeric structure of PSI on longer time scales. We find that docking of the soluble electron mediators cytochrome c6 and ferredoxin is not impacted by the presence of detergent, differing only slightly for predictions based on our MD model of detergent-solubilized PSI relative to predictions based on the PSI crystal structure. We attribute the slight changes in binding to fluctuations in binding site residues during the simulation, which we believe may imply an inducedfit mechanism for mediator docking. These findings yield new insights into the structural integrity and activity of this protein in vitro. In conclusion, the molecular dynamics simulation and analyses reported in this paper are a significant step in understanding the solution structure and dynamics of detergent-solubilized membrane proteins at the atomic level. Furthermore, the results reported here in particular offer a solid starting point for understanding the in vitro structure and dynamics of the photoactive pigment−protein complex known as photosystem I, a protein that functions as a key component of oxygenic photosynthesis in plants and microorganisms. Of particular significance is the indication that the inclusion of associated lipids may be important for maintaining the stability of trimeric PSI in detergent solution. Additionally, the presence of DDM detergent does not appear to interfere with PSI activity. This information may prove useful for prolonging the stability and activity of PSI in vitro in order to improve photosynthesis-based energy conversion devices.

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CONCLUSIONS We have performed MD simulations of the detergentsolubilized PSI trimer, one of the largest membrane protein complexes known to have so far been studied. RMSD analysis has shown it to be a stable complex, with comparison of Cα RMSD values suggesting that the behavior of the protein in a DDM micellar environment is similar to that observed for previous MD studies of membrane proteins in lipid bilayers. Furthermore, we have found that the largest contributors to the structural drift of this protein compared to its crystal structure are the peripheral transmembrane subunits, which are in close contact with detergent and in general become more compact and kinked over the course of the simulation. We also investigated protein local mobility based on RMSF, concluding that regions of low local mobility were confined to the core residues of the reaction center that are not solvent-exposed, with regions of high fluctuation corresponding to turns in the transmembrane α-helices and flexible loops of the stromal domains, all of which are exposed to the solvent. We examined differences in detergent behavior in our protein−detergent complex versus a pure detergent micelle. In terms of micelle geometry, we find that the thickness of the detergent belt shrinks over the course of MD simulation as it conforms to the nonuniform hydrophobic periphery of the protein, with the complex as a whole becoming more ellipsoidal in shape as a result; a similar ellipsoidal shape was observed for MD simulation of the pure detergent micelle. Examining radial density distributions for various system components in each case, we find that the presence of protein results in a broadening of the detergent head and tail atom density profiles but has little effect on the extent of water penetration. We further note that the distance between the peaks of the head and tail atom density curves decreases from 28 Å for the pure DDM micelle to 19 Å for the PSI−DDM complex, suggesting that favorable interactions with the hydrophobic transmembrane domain of PSI result in the shrinking of the detergent molecules. Calculation of solvent-exposed surface area (SASA) reveals DDM is overall less densely packed in the presence of protein compared to a pure detergent micelle but with differing trends in the head and tail groups. We have further shown that the detergent tails are more ordered in the PSI−DDM complex compared to a pure DDM micelle, based on evaluation of the dihedral angle order parameters (S2) of the detergent tail CH2 groups and MSD values for the associated hydrogen atoms. Taken in its entirety, we believe the behavior of the transmembrane domain of PSI and the surrounding detergent molecules suggest a degree of plasticity in the structure of the in vivo complex. Through kinking and tilting motions of the peripheral transmembrane helices and the rearrangement of surrounding lipid molecules, the membrane-shielded region of the protein can adapt to the lipid bilayer thickness and vice versa, thus allowing PSI to adapt to different lipid bilayer environments (i.e., different lipid types or phases). We have also attempted to discern any functional consequences for this photoactive pigment−protein complex due to solution dynamics and protein−detergent interactions. We attribute the extension of the N-terminus of psaL from the trimer voids out into the solvent during the course of MD simulation to the absence of integral lipids found in the crystal structure of PSI from our simulation, and believe that it could

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ASSOCIATED CONTENT

* Supporting Information S

Figures showing a plot of Cα RMSD versus time, radial atom density distributions, and detergent solvent-exposed surface area versus time. Table showing radial distances corresponding to maximum atomic densities of detergent head and tail atoms. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*Address: 419 Dougherty Engineering Building, Department of Chemical and Biomolecular Engineering, University of Tennessee, Knoxville, TN 37996. E-mail: [email protected]. Author Contributions

The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors would like to acknowledge financial support for B.J.H. from the NSF-EPSCoR sponsored TN-SCORE (NSF EPS-1004083). This material is based upon work performed on the Cray XE6 supercomputer Hopper, a resource of the National Energy Research Scientific Computing Center, which is supported by the Office of Science of the U.S. Department of Energy under Contract No. DE-AC02-05CH11231.

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ABBREVIATIONS chl a, chlorophyll a; cyt c6, cytochrome c6; DDM, n-dodecyl-βmaltoside; DMPC, dimyristoyl phosphatidylcholine; DPC, dodecylphosphocholine; Fd, ferredoxin; GpA, glycophorin A; 11642



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(19) Kiley, P.; Zhao, X. J.; Vaughn, M.; Baldo, M. A.; Bruce, B. D.; Zhang, S. G. Self-Assembling Peptide Detergents Stabilize Isolated Photosystem I on a Dry Surface for an Extended Time. PLoS Biol. 2005, 3, 1180−1186. (20) Nelson, M. T.; Humphrey, W.; Gursoy, A.; Dalke, A.; Kale, L. V.; Skeel, R. D.; Schulten, K. NAMD: A Parallel, Object Oriented Molecular Dynamics Program. Int. J. Supercomput. Appl. High Perform. Comput. 1996, 10, 251−268. (21) Lindahl, E.; Hess, B.; van der Spoel, D. Gromacs 3.0: A Package for Molecular Simulation and Trajectory Analysis. J. Mol. Model. 2001, 7, 306−317. (22) Case, D. A.; Cheatham, T. E.; Darden, T.; Gohlke, H.; Luo, R.; Merz, K. M.; Onufriev, A.; Simmerling, C.; Wang, B.; Woods, R. J. The Amber Biomolecular Simulation Programs. J. Comput. Chem. 2005, 26, 1668−1688. (23) Hansson, T.; Oostenbrink, C.; van Gunsteren, W. F. Molecular Dynamics Simulations. Curr. Opin. Struct. Biol. 2002, 12, 190−196. (24) Venable, R. M.; Zhang, Y. H.; Hardy, B. J.; Pastor, R. W. Molecular-Dynamics Simulations of a Lipid Bilayer and of Hexadecane - an Investigation of Membrane Fluidity. Science 1993, 262, 223−226. (25) Egberts, E.; Marrink, S. J.; Berendsen, H. J. C. MolecularDynamics Simulation of a Phospholipid Membrane. Eur. Biophys. J. Biophys. Lett. 1994, 22, 423−436. (26) Feller, S. E.; Pastor, R. W. Constant Surface Tension Simulations of Lipid Bilayers: The Sensitivity of Surface Areas and Compressibilities. J. Chem. Phys. 1999, 111, 1281−1287. (27) Tieleman, D. P.; Berendsen, H. J. C. A Molecular Dynamics Study of the Pores Formed by Escherichia Coli OmpF Porin in a Fully Hydrated Palmitoyloleoylphosphatidylcholine Bilayer. Biophys. J. 1998, 74, 2786−2801. (28) Im, W.; Roux, B. Ions and Counterions in a Biological Channel: A Molecular Dynamics Simulation of OmpF Porin from Escherichia Coli in an Explicit Membrane with 1 M KCl Aqueous Salt Solution. J. Mol. Biol. 2002, 319, 1177−1197. (29) Bond, P. J.; Faraldo-Gomez, J. D.; Sansom, M. S. P. OmpA: A Pore or Not a Pore? Simulation and Modeling Studies. Biophys. J. 2002, 83, 763−775. (30) de Groot, B. L.; Grubmuller, H. Water Permeation across Biological Membranes: Mechanism and Dynamics of Aquaporin-1 and Glpf. Science 2001, 294, 2353−2357. (31) Zhu, F. Q.; Tajkhorshid, E.; Schulten, K. Molecular Dynamics Study of Aquaporin-1 Water Channel in a Lipid Bilayer. FEBS Lett. 2001, 504, 212−218. (32) Woolf, T. B.; Roux, B. Structure, Energetics, and Dynamics of Lipid-Protein Interactions: A Molecular Dynamics Study of the Gramicidin A Channel in a DMPC Bilayer. Proteins: Struct., Funct., Genet. 1996, 24, 92−114. (33) Chiu, S. W.; Subramaniam, S.; Jakobsson, E. Simulation Study of a Gramicidin/Lipid Bilayer System in Excess Water and Lipid. I. Structure of the Molecular Complex. Biophys. J. 1999, 76, 1929−1938. (34) Berneche, S.; Roux, B. Molecular Dynamics of the KcsA K+ Channel in a Bilayer Membrane. Biophys. J. 2000, 78, 2900−2917. (35) Shrivastava, I. H.; Sansom, M. S. P. Simulations of Ion Permeation through a Potassium Channel: Molecular Dynamics of KcsA in a Phospholipid Bilayer. Biophys. J. 2000, 78, 557−570. (36) Elmore, D. E.; Dougherty, D. A. Molecular Dynamics Simulations of Wild-Type and Mutant Forms of the Mycobacterium Tuberculosis MscL Channel. Biophys. J. 2001, 81, 1345−1359. (37) Gullingsrud, J.; Kosztin, D.; Schulten, K. Structural Determinants of MscL Gating Studied by Molecular Dynamics Simulations. Biophys. J. 2001, 80, 2074−2081. (38) Huber, T.; Botelho, A. V.; Beyer, K.; Brown, M. F. Membrane Model for the G-Protein-Coupled Receptor Rhodopsin: Hydrophobic Interface and Dynamical Structure. Biophys. J. 2004, 86, 2078−2100. (39) Periole, X.; Huber, T.; Marrink, S. J.; Sakmar, T. P. G ProteinCoupled Receptors Self-Assemble in Dynamics Simulations of Model Bilayers. J. Am. Chem. Soc. 2007, 129, 10126−10132. (40) Nygaard, R.; Zou, Y. Z.; Dror, R. O.; Mildorf, T. J.; Arlow, D. H.; Manglik, A.; Pan, A. C.; Liu, C. W.; Fung, J. J.; Bokoch, M. P.; et al.

GPCR, G-protein-coupled receptor; LHCII, light-harvesting complex II; LHG, 1,2-dipalmitoyl-phosphatidylglycerol; LMG, 1,2-distearoyl-monogalactosyldiglyceride; MSD, mean-square displacement; MD, molecular dynamics; OES, N-octyl-2hydroxyethyl sulfoxide; POPC, palmitoyl oleoyl phosphatidylcholine; PSI, photosystem I; PSII, photosystem II; RMSD, root-mean-square displacement; RMSF, root-mean-square fluctuation; SANS, small-angle neutron scattering; SASA, solvent-accessible surface area; T. elongatus, Thermosynechococcus elongatus

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REFERENCES

(1) Wallin, E.; von Heijne, G. Genome-Wide Analysis of Integral Membrane Proteins from Eubacterial, Archaean, and Eukaryotic Organisms. Protein Sci. 1998, 7, 1029−1038. (2) White, S. H. Biophysical Dissection of Membrane Proteins. Nature 2009, 459, 344−346. (3) White, S. H.; Wimley, W. C. Membrane Protein Folding and Stability: Physical Principles. Annu. Rev. Biophys. Biomol. Struct. 1999, 28, 319−365. (4) Garavito, R. M.; Ferguson-Miller, S. Detergents as Tools in Membrane Biochemistry. J. Biol. Chem. 2001, 276, 32403−32406. (5) Tamm, L. K.; Liang, B. Y. NMR of Membrane Proteins in Solution. Prog. Nucl. Magn. Reson. Spectrosc. 2006, 48, 201−210. (6) Lipfert, J.; Doniach, S. Small-Angle X-Ray Scattering from RNA, Proteins, and Protein Complexes. Annu. Rev. Biophys. Biomol. Struct. 2007, 36, 307−327. (7) Jordan, P.; Fromme, P.; Witt, H. T.; Klukas, O.; Saenger, W.; Krauss, N. Three-Dimensional Structure of Cyanobacterial Photosystem I at 2.5 Angstrom Resolution. Nature 2001, 411, 909−917. (8) Golbeck, J. H. Structure and Function of Photosystem-I. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1992, 43, 293−324. (9) Greenbaum, E. Platinized Chloroplasts - a Novel Photocatalytic Material. Science 1985, 230, 1373−1375. (10) Iwuchukwu, I. J.; Vaughn, M.; Myers, N.; O’Neill, H.; Frymier, P.; Bruce, B. D. Self-Organized Photosynthetic Nanoparticle for CellFree Hydrogen Production. Nat. Nanotechnol. 2010, 5, 73−79. (11) Iwuchukwu, I. J.; Iwuchukwu, E.; Le, R.; Paquet, C.; Sawhney, R.; Bruce, B.; Frymier, P. Optimization of Photosynthetic Hydrogen Yield from Platinized Photosystem I Complexes Using Response Surface Methodology. Int. J. Hydrogen Energy 2011, 36, 11684−11692. (12) Schwarze, A.; Kopczak, M. J.; Rogner, M.; Lenz, O. Requirements for Construction of a Functional Hybrid Complex of Photosystem I and [NiFe]-Hydrogenase. Appl. Environ. Microbiol. 2010, 76, 2641−2651. (13) Lubner, C. E.; Applegate, A. M.; Knorzer, P.; Ganago, A.; Bryant, D. A.; Happe, T.; Golbeck, J. H. Solar Hydrogen-Producing Bionanodevice Outperforms Natural Photosynthesis. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 20988−20991. (14) Lee, I.; Lee, J. W.; Greenbaum, E. Biomolecular Electronics: Vectorial Arrays of Photosynthetic Reaction Centers. Phys. Rev. Lett. 1997, 79, 3294−3297. (15) Das, R.; Kiley, P. J.; Segal, M.; Norville, J.; Yu, A. A.; Wang, L. Y.; Trammell, S. A.; Reddick, L. E.; Kumar, R.; Stellacci, F.; et al. Integration of Photosynthetic Protein Molecular Complexes in SolidState Electronic Devices. Nano Lett. 2004, 4, 1079−1083. (16) Ciesielski, P. N.; Faulkner, C. J.; Irwin, M. T.; Gregory, J. M.; Tolk, N. H.; Cliffel, D. E.; Jennings, G. K. Enhanced Photocurrent Production by Photosystem I Multilayer Assemblies. Adv. Funct. Mater. 2010, 20, 4048−4054. (17) LeBlanc, G.; Chen, G. P.; Gizzie, E. A.; Jennings, G. K.; Cliffel, D. E. Enhanced Photocurrents of Photosystem I Films on p-Doped Silicon. Adv. Mater. 2012, 24, 5959−5962. (18) Mershin, A.; Matsumoto, K.; Kaiser, L.; Yu, D. Y.; Vaughn, M.; Nazeeruddin, M. K.; Bruce, B. D.; Graetzel, M.; Zhang, S. G. SelfAssembled Photosystem-I Biophotovoltaics on Nanostructured TiO2 and ZnO. Sci. Rep. 2012, 2, 1−7. 11643



Article

and Condensed Phase Macromolecular Target Data. J. Comput. Chem. 2000, 21, 86−104. (61) Zhang, L.; Silva, D. A.; Yan, Y. J.; Huang, X. H. Force Field Development for Cofactors in the Photosystem II. J. Comput. Chem. 2012, 33, 1969−1980. (62) Smith, D. M. A.; Xiong, Y. J.; Straatsma, T. P.; Rosso, K. M.; Squier, T. C. Force-Field Development and Molecular Dynamics of NiFe Hydrogenase. J. Chem. Theory Comput. 2012, 8, 2103−2114. (63) Darden, T.; York, D.; Pedersen, L. Particle Mesh Ewald - an N.Log(N) Method for Ewald Sums in Large Systems. J. Chem. Phys. 1993, 98, 10089−10092. (64) Feller, S. E.; Zhang, Y. H.; Pastor, R. W.; Brooks, B. R. Constant-Pressure Molecular-Dynamics Simulation - the Langevin Piston Method. J. Chem. Phys. 1995, 103, 4613−4621. (65) Humphrey, W.; Dalke, A.; Schulten, K. VMD: Visual Molecular Dynamics. J. Mol. Graphics Modell. 1996, 14, 33−38. (66) Pierce, B. G.; Wiehe, K.; Hwang, H.; Kim, B.-H.; Vreven, T.; Weng, Z. ZDOCK Server: Interactive Docking Prediction of ProteinProtein Complexes and Symmetric Multimers. Bioinformatics 2014, 30, 1771−1773. (67) Sommer, F.; Drepper, F.; Haehnel, W.; Hippler, M. The Hydrophobic Recognition Site Formed by Residues PsaA-Trp(651) and PsaB-Trp(627) of Photosystem I in Chlamydomonas Reinhardtii Confers Distinct Selectivity for Binding of Plastocyanin and Cytochrome c(6). J. Biol. Chem. 2004, 279, 20009−20017. (68) Setif, P.; Fischer, N.; Lagoutte, B.; Bottin, H.; Rochaix, J. D. The Ferredoxin Docking Site of Photosystem I. Biochim. Biophys. Acta, Bioenerg. 2002, 1555, 204−209. (69) Mukherjee, D.; May, M.; Khomami, B. Detergent-Protein Interactions in Aqueous Buffer Suspensions of Photosystem I (PSI). J. Colloid Interface Sci. 2011, 358, 477−484. (70) Bond, P. J.; Sansom, M. S. P. Membrane Protein Dynamics Versus Environment: Simulations of OmpA in a Micelle and in a Bilayer. J. Mol. Biol. 2003, 329, 1035−1053. (71) Deol, S. S.; Bond, P. J.; Domene, C.; Sansom, M. S. P. LipidProtein Interactions of Integral Membrane Proteins: A Comparative Simulation Study. Biophys. J. 2004, 87, 3737−3749. (72) Zilber, A. L.; Malkin, R. Organization and Topology of Photosystem-I Subunits. Plant Physiol. 1992, 99, 901−911. (73) Psachoulia, E.; Bond, P. J.; Sansom, M. S. P. MD Simulations of Mistic: Conformational Stability in Detergent Micelles and Water. Biochemistry 2006, 45, 9053−9058. (74) Aponte-Santamaria, C.; Briones, R.; Schenk, A. D.; Walz, T.; de Groot, B. L. Molecular Driving Forces Defining Lipid Positions around Aquaporin-0. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 9887−9892. (75) vanderSpoel, D.; Berendsen, H. J. C. Molecular Dynamics Simulations of Leu-Enkephalin in Water and DMSO. Biophys. J. 1997, 72, 2032−2041. (76) Edidin, M. The State of Lipid Rafts: From Model Membranes to Cells. Annu. Rev. Biophys. Biomol. Struct. 2003, 32, 257−283. (77) Simons, K.; Vaz, W. L. C. Model Systems, Lipid Rafts, and Cell Membranes. Annu. Rev. Biophys. Biomol. Struct. 2004, 33, 269−295. (78) Kouril, R.; Zygadlo, A.; Arteni, A. A.; de Wit, C. D.; Dekker, J. P.; Jensen, P. E.; Scheller, H. V.; Boekema, E. J. Structural Characterization of a Complex of Photosystem I and Light-Harvesting Complex II of Arabidopsis Thaliana. Biochemistry 2005, 44, 10935− 10940. (79) Amunts, A.; Nelson, N. Plant Photosystem I Design in the Light of Evolution. Structure 2009, 17, 637−650. (80) Fromme, P.; Melkozernov, A.; Jordan, P.; Krauss, N. Structure and Function of Photosystem I: Interaction with Its Soluble Electron Carriers and External Antenna Systems. FEBS Lett. 2003, 555, 40−44. (81) Lange, C.; Nett, J. H.; Trumpower, B. L.; Hunte, C. Specific Roles of Protein-Phospholipid Interactions in the Yeast Cytochrome bc(1) Complex Structure. EMBO J. 2001, 20, 6591−6600. (82) Fyfe, P. K.; Isaacs, N. W.; Cogdell, R. J.; Jones, M. R. Disruption of a Specific Molecular Interaction with a Bound Lipid Affects the Thermal Stability of the Purple Bacterial Reaction Centre. Biochim. Biophys. Acta, Bioenerg. 2004, 1608, 11−22.

The Dynamic Process of Beta(2)-Adrenergic Receptor Activation. Cell 2013, 152, 532−542. (41) Chandler, D. E.; Hsin, J.; Harrison, C. B.; Gumbart, J.; Schulten, K. Intrinsic Curvature Properties of Photosynthetic Proteins in Chromatophores. Biophys. J. 2008, 95, 2822−2836. (42) Sener, M.; Strumpfer, J.; Timney, J. A.; Freiberg, A.; Hunter, C. N.; Schulten, K. Photosynthetic Vesicle Architecture and Constraints on Efficient Energy Harvesting. Biophys. J. 2010, 99, 67−75. (43) Chandler, D. E.; Strumpfer, J.; Sener, M.; Scheuring, S.; Schulten, K. Light Harvesting by Lamellar Chromatophores in Rhodospirillum Photometricum. Biophys. J. 2014, 106, 2503−2510. (44) 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, 15670−15673. (45) Tarek, M.; Bandyopadhyay, S.; Klein, M. L. Molecular Dynamics Studies of Aqueous Surfactants Systems. J. Mol. Liq. 1998, 78, 1−6. (46) Bruce, C. D.; Berkowitz, M. L.; Perera, L.; Forbes, M. D. E. Molecular Dynamics Simulation of Sodium Dodecyl Sulfate Micelle in Water: Micellar Structural Characteristics and Counterion Distribution. J. Phys. Chem. B 2002, 106, 3788−3793. (47) Bocker, J.; Brickmann, J.; Bopp, P. Molecular-Dynamics Simulation Study of an N-Decyltrimethylammonium Chloride Micelle in Water. J. Phys. Chem. 1994, 98, 712−717. (48) Bogusz, S.; Venable, R. M.; Pastor, R. W. Molecular Dynamics Simulations of Octyl Glucoside Micelles: Dynamic Properties. J. Phys. Chem. B 2001, 105, 8312−8321. (49) Abel, S.; Dupradeau, F. Y.; Raman, E. P.; MacKerell, A. D.; Marchi, M. Molecular Simulations of Dodecyl-Beta-Maltoside Micelles in Water: Influence of the Headgroup Conformation and Force Field Parameters. J. Phys. Chem. B 2011, 115, 487−499. (50) Wendoloski, J. J.; Kimatian, S. J.; Schutt, C. E.; Salemme, F. R. Molecular-Dynamics Simulation of a Phospholipid Micelle. Science 1989, 243, 636−638. (51) Wymore, T.; Gao, X. F.; Wong, T. C. Molecular Dynamics Simulation of the Structure and Dynamics of a Dodecylphosphocholine Micelle in Aqueous Solution. J. Mol. Struct. 1999, 485, 195−210. (52) Abel, S.; Dupradeau, F.-Y.; Marchi, M. Molecular Dynamics Simulations of a Characteristic DPC Micelle in Water. J. Chem. Theory Comput. 2012, 8, 4610−4623. (53) Braun, R.; Engelman, D. M.; Schulten, K. Molecular Dynamics Simulations of Micelle Formation around Dimeric Glycophorin A Transmembrane Helices. Biophys. J. 2004, 87, 754−763. (54) Bond, P. J.; Cuthbertson, J. M.; Deol, S. S.; Sansom, M. S. P. MD Simulations of Spontaneous Membrane Protein/Detergent Micelle Formation. J. Am. Chem. Soc. 2004, 126, 15948−15949. (55) Rodriguez-Ropero, F.; Fioroni, M. Structural and Dynamical Analysis of an Engineered FhuA Channel Protein Embedded into a Lipid Bilayer or a Detergent Belt. J. Struct. Biol. 2012, 177, 291−301. (56) Koutsioubas, A.; Berthaud, A.; Mangenot, S.; Perez, J. Ab Initio and All-Atom Modeling of Detergent Organization around Aquaporin0 Based on SAXS Data. J. Phys. Chem. B 2013, 117, 13588−13594. (57) Le, R. K.; Harris, B. J.; Iwuchukwu, I. J.; Bruce, B. D.; Cheng, X.; Qian, S.; Heller, W. T.; O’Neill, H.; Frymier, P. D. Analysis of the Solution Structure of Thermosynechococcus Elongatus Photosystem I in N-Dodecyl-Beta-D-Maltoside Using Small-Angle Neutron Scattering and Molecular Dynamics Simulation. Arch. Biochem. Biophys. 2014, 550−551, 50−57. (58) Martinez, L.; Andrade, R.; Birgin, E. G.; Martinez, J. M. Packmol: A Package for Building Initial Configurations for Molecular Dynamics Simulations. J. Comput. Chem. 2009, 30, 2157−2164. (59) Klauda, J. B.; Venable, R. M.; Freites, J. A.; O’Connor, J. W.; Tobias, D. J.; Mondragon-Ramirez, C.; Vorobyov, I.; MacKerell, A. D., Jr.; Pastor, R. W. Update of the CHARMM All-Atom Additive Force Field for Lipids: Validation on Six Lipid Types. J. Phys. Chem. B 2010, 114, 7830−7843. (60) Foloppe, N.; MacKerell, A. D. All-Atom Empirical Force Field for Nucleic Acids: I. Parameter Optimization Based on Small Molecule 11644



Article

(83) Pendley, S. S.; Manocchi, A. K.; Baker, D. R.; Sumner, J. J.; Lundgren, C. A.; Hurley, M. M. Challenges and Development of a Multi-Scale Computational Model for Photosystem I Decoupled Energy Conversion. In Applications of Molecular Modeling to Challenges in Clean Energy; Fitzgerald, G., Govind, N., Eds.; 2013; Vol. 1133, pp 177−202. (84) Fischer, N.; Setif, P.; Rochaix, J. D. Site-Directed Mutagenesis of the PsaC Subunit of Photosystem I - F-B Is the Cluster Interacting with Soluble Ferredoxin. J. Biol. Chem. 1999, 274, 23333−23340. (85) Sommer, F.; Drepper, F.; Hippler, M. The Luminal Helix L of PsaB Is Essential for Recognition of Plastocyanin or Cytochrome c(6) and Fast Electron Transfer to Photosystem I in Chlamydomonas Reinhardtii. J. Biol. Chem. 2002, 277, 6573−6581.

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