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Jan 16, 2017 - Molecular Details of the PH Domain of ACAP1BAR‑PH Protein Binding to PIP-Containing Membrane. Kevin Chun Chan,. †. Lanyuan Lu,. ‡...
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Molecular Details of the PH Domain of ACAP1BAR‑PH Protein Binding to PIP-Containing Membrane Kevin Chun Chan,† Lanyuan Lu,‡ Fei Sun,§,∥ and Jun Fan*,†,⊥,# †

Department of Physics and Materials Science, City University of Hong Kong, Hong Kong, China School of Biological Sciences, Nanyang Technological University, 639798, Singapore § National Laboratory of Biomacromolecules, Institute of Biophysics, Chinese Academy of Sciences, Beijing 100101, China ∥ Center for Biological Imaging, Institute of Biophysics, Chinese Academy of Sciences, Beijing 100101, China ⊥ City University of Hong Kong Shenzhen Research Institute, Shenzhen 518057, China # Center for Advanced Nuclear Safety and Sustainable Development, City University of Hong Kong, Hong Kong, China ‡

S Supporting Information *

ABSTRACT: ACAP1 proteins were previously reported to specifically bind PIP2-containing cell membranes and form well-structured protein lattices in order to conduct membrane tubulation. We carried out molecular dynamics simulations to characterize orientation of the PH domains with respect to the BAR domains inside the protein dimer. Followed by molecular dynamics simulations, we present a comprehensive orientation analysis of PH domain under different states including unbound and bound with lipids. We have examined two binding pockets on the PH domain and present PMF profiles of the two pockets to account for their preference to PIP2 lipids. Combining orientation analysis and studies of binding pockets, our simulations results reveal valuable molecular basis for protein−lipid interactions of ACAP1 proteins during membrane remodeling process.

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INTRODUCTION Understanding the association of peripheral proteins to cell membranes is critical for many aspects of cellular function.1,2 One of the fundamental questions would be how proteins recognize and bind to specific lipid molecules.3−6 Addressing this question requires knowledge of structure, energetics, and dynamics of the complex protein−lipid systems. Molecular dynamics (MD) simulations have been widely employed as one of the best tools for studying such complex biological systems on both temporal and spatial scales not accessible to current experimental methods.7−12 MD simulations provide not only molecular details of the structures and dynamics of the protein−lipid complexes, but also valuable energetics information when combining with sophisticated free-energy calculation methodologies.13−18 Arfgap with coiled coil, Ankyrin repeat, and PH domain protein 1 (ACAP1) were initially characterized as a GTPaseactivating protein (GAP) for ADP-ribosylation factor 6 (ARF6).19 Subsequently, it was found to also involve in endocytic recycling as an ARF6 effector.20,21 Being a coat component, four domains of ACAP1 were found to be responsible for different functions during endocytosis: ArfGAP and ankryin repeat (ANK) domains have been well studied for its cargo binding abilities;22 Bin-Amphiphysin-Rvs (BAR) and Pleckstrin homology (PH) domains were found to oligomerize and promote membrane curvatures in order to generate © 2017 American Chemical Society

transport carriers. BAR domain is one of the best studied protein domain motif related to membrane curvature recognition and generation.23−31 In previous studies, however, we found that the PH domain of ACAP1 protein dimer take over the role of the BAR domain when binding to cell membranes.32 The PH domain was first identified in 1993 and has been found in numerous proteins involved in cellular signaling.33,34 The view that the PH domain directs membrane targeting of their host proteins by binding to phosphoinositides (PIs) has been established since then.35−37 PIs play crucial roles in both cell signaling and membrane traffic.38−41 Phosphatidylinositol4,5-bisphosphate (PI(4,5)P2 or more briefly PIP2), being the best studied species among eukaryotic PIs, were reported to be bound by a significant number of known PH domains with high affinity and specificity.42,43 In addition to PIs, PH domains were also reported to bind to other proteins in signal transduction pathways.44,45 Multiple functions of the PH domain revealed by experiments initiate a number of computational studies.9−12,46 While Special Issue: Klaus Schulten Memorial Issue Received: September 21, 2016 Revised: January 15, 2017 Published: January 16, 2017 3586

DOI: 10.1021/acs.jpcb.6b09563 J. Phys. Chem. B 2017, 121, 3586−3596

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energy, temperature, pressure, and root-mean-square deviation (RMSD) of the protein backbone. All the quantities reached a plateau after equilibrium, while the RMSD profile being the latest one (Figure S1). Only data after the system was equilibrated were taken for further analysis. For the protein− solvent systems, the first 180 ns of trajectories were excluded for analysis. For the 420 and 250 ns protein−lipid system, the first 220 and 150 ns of trajectories were excluded for analysis. All quantities presented in this article are averaged values over the equilibrated simulation windows. Protein−lipid contacts were defined as a cutoff distance of 12 Å between the center of mass (COM) of a residue and any lipid backbone phosphate. Such distance was chosen according to the cutoff distance of the nonbonded interactions. Umbrella Sampling Simulations. The PH domain of the ACAP1BAR‑PH structure from the last frame of the protein−lipid simulation 1 served as the initial PIP2-bound structure for umbrella sampling. As we have PIP2 lipid found at both binding pockets on the PH domain, we generated two structures, in which PIP2 lipid bound to only one of the pockets, by replacing one of the PIP2 with a DOPC molecule. We then placed the new PIP2-bound structure inside DOPC lipid bilayers, so that the bound PIP2 is the only PIP2 lipid in the membrane. In this way, we may remove the effects of surrounding charged lipids. POPS-bound structures were prepared in a similar manner by replacing the PIP2 lipid inside the desired pocket into a POPS lipid. Using a steered molecular dynamics (SMD) simulation, the PH domain was pulled away from the bound PIP2 lipid by fixing the COM to a reference point (with restraint constant k = 1000 kJ mol−1 nm−2) which was moved along the membrane normal at a rate of 1 Å ns−1 for a total distance of 18 Å. In a SMD simulation, the orientation of the PH domain and the PIP2 backbone phosphate atom was restrained relative to their initial orientation or position using collective variable-based calculations. Snapshots at separation intervals of 1.2 Å were used as initial configurations to define a reaction coordinate for 16 windows of umbrella sampling (US) simulations. The first window was the initial bound state of PH domain. The first 11 windows were let to relax for 100 ns, followed by 50 ns of productive simulation; while the last 5 windows were relaxed for 50 ns, followed by 50 ns of productive simulation (the total amount of simulation per system was 2150 ns). As a result, window centers ranging from 58.3 to 70.3 Å were used for the free energy calculations. Further extended windows (71.5 Å to 76.3 Å) were added for orientation analysis of binding pockets. To eliminate perturbation to the PIP2 lipid by US window restraints, we have chosen the separation of protein COM and lower-layer lipid backbone phosphorus atoms COM to be restrained as the window centers (with restraint constant k = 1000 kJ mol−1 nm−2). No further restraints were added to the system, except the US window restraints. Potential of mean force (PMF) profiles were calculated using the GROMACS g_wham tool.58 Convergence was assessed by calculating profiles for 30 ns intervals in 10 ns sliding blocks (Figure S2). Data before equilibrium were discarded in order to calculate the final profile. Errors were obtained from bootstrap analysis using 200 bootstraps. All profiles have been shifted so that the PMF value in bulk is 0 kcal mol−1.

most of the previous studies were restricted to the PH domain of GRP1 protein, we recently performed experimental and computational studies on BAR-PH domain of ACAP1 protein (ACAP1BAR‑PH) to address novel features of the PH domain overtaking the role of the BAR domain during membrane curvature generation.32 In current studies, we further investigate the molecular details of the protein−lipid interactions of ACAP1BAR‑PH protein with the assistance of both restrained and unbiased atomistic molecular dynamics (MD) simulations and sophisticated free-energy calculation methodologies.

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METHODS Molecular Dynamics Simulations. A single ACAP1BAR‑PH dimer with and without lipid bilayers underneath were constructed in a similar manner. Initial structures of the protein were based on the crystal structure of ACAP1BAR‑PH protein (PDB: 4NSW). The lipid bilayer was constructed using CHARMM-GUI47 with a composition of DOPC/DOPE/ POPS/PI(4,5)P2 (4:3:2:1) and contained 600 lipid molecules in total. Details of the membrane construction can be referred to our previous work.32 The protein was placed on the top of the membrane for the protein−membrane system. Orientations of the protein were chosen so that the concave surface of the BAR domain faces the underneath membrane. The distance between the protein and the membrane was chosen to be 10 A, within the cutoff of the nonbonded interactions. The systems were then solvated in explicit TIP3P water molecules,48 with potassium and chloride ions included at a final concentration of 0.18 M using the solvate and autoionize tools in VMD.49 Periodic boundary conditions were introduced to both systems. Overall, three protein−solvent simulations (210 × 100 × 110 Å3, ∼ 220k atoms, simulation 1−3) and two protein−lipid simulations (270 × 105 × 155 Å3, ∼ 300k atoms, simulation 4− 5) have been performed. The MD simulations were performed using NAMD 2.11 package.50 CHARMM 22/27 force field51 with CMAP correction52,53 was used for proteins and CHARMM 36 force field54 was used for lipids. Electrostatic interactions were calculated using the particle mesh Ewald sum method55 with a cutoff of 12 Å. All hydrogen-containing covalent bonds were constrained by the SHAKE algorithm,56 therefore allowing an integration time step of 2 fs. Initially, the system was minimized using conjugate gradient algorithm followed by an equilibrium in canonical ensemble at 310 K. Protein backbone and oxygen atoms of water were harmonically restrained by a spring constant (1 kcal mol−1 Å−2). Simulations were then continued in the constant NPT ensemble (310 K and 1 atm). Langevin thermostats with a damping coefficient of 0.5 ps−1 were used to control the system temperature. A Langevin-piston barostat57 with a piston period of 2 ps and a damping time of 2 ps was used to control the pressure. Anisotropic fluctuations were allowed for the protein−lipid systems. Constraints were next released stepwise (with spring constant gradually decrease from 1 kcal mol−1 Å2 to 0 by steps of 0.1 kcal mol−1 Å2) before starting the production runs. A total of 420 ns and 2 × 380 ns of trajectory were generated for the protein−solvent systems, respectively. A total of 420 and 250 ns of trajectories were generated for the two protein− lipid systems, respectively. In both simulations, we have observed that only one of the PH domains bound to the membrane underneath, i.e., the asymmetrical binding event. Equilibrium of the systems were judged by examining a number of thermodynamic quantities, such as simulation box size,

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RESULTS MD simulations grant us valuable molecular details of dynamical biological systems. In order to study interactions between ACAP1BAR‑PH protein dimer and PIP2-containing 3587

DOI: 10.1021/acs.jpcb.6b09563 J. Phys. Chem. B 2017, 121, 3586−3596

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The Journal of Physical Chemistry B multicomponent phospholipid membranes, we have performed a number of atomistic MD simulations, including (1) one single ACAP1BAR‑PH protein dimer in ionic solvent and (2) one single ACAP1BAR‑PH protein dimer interacting with PIP2-containing bilayers in ionic solvent. Combining multiple atomistic MD simulations not only improved statistics of analysis by reducing errors due to insufficient sampling, but also uncovered interesting protein dynamics under different environments. Within the availability of computational resources, all molecular systems have been performed multiple times of simulations for ensuring repeatability and improving analytical statistics (simulation details in Methods Section). Hinted by the molecular details of interactions between ACAP1BAR‑PH protein dimer with PIP2-containing bilayers, we also identified two binding pockets on the PH domain of ACAP1BAR‑PH protein dimer for PIP2 lipids. We found that although both pockets bound to PIP2 lipids consistently in multiple atomistic MD simulations, the binding pockets could indeed behave very differently. In addition to ordinary unbiased MD simulations, multiple restrained MD simulations have also been performed as umbrella sampling (US) in order to compute the potential of mean force (PMF) profiles. One single PH domain was obtained from the ACAP1BAR‑PH protein dimer and placed above either a PIP2- or POPS-containing bilayers. As a result, PMF profiles of a PH domain interacting with a PIP2 or POPS lipid molecule along the normal of the membrane were obtained (simulation details in Methods Section). We have performed four US simulations in total in which the two binding pockets were bound to either PIP2 or POPS lipid, respectively. Combining unbiased and restrained MD simulations, we examined carefully the nature of the two binding pockets. Flexible Loop Linking the PH Domain and the BAR Domain. The BAR and PH domains were reported, respectively, as conserved motifs in many different proteins.23,33−35 In order to study the cooperative work carried out by these two domains upon the membrane binding events, we sought to examine the relative motion of the PH domain with respect to the BAR domain in ACAP1 proteins. By computing the three principle axes of the BAR domain, we obtained the orientation of the BAR domain as the three reference axes (yellow arrows in Figure 1). Orientation of the PH domain was defined by one of the principle axes of the sole helix (residue 345−361) in the PH domain pointing along the

length of the helix (blue arrows in Figure 1). As an ACAP1BAR‑PH domain is a dimeric structure of two BAR-PH monomers, the structure shows 2-fold symmetry. Therefore, when computing the three reference axes for each PH domain in an ACAP1BAR‑PH protein dimer, we would invert the first and the third principle axes of the BAR domain. As a result, the first principle axis of the BAR domain always points in the direction of the blue arrow of the desired PH domain; the third principle axis of the BAR domain always points from the geometry center of the BAR domain toward the desired PH domain; while the second principle axis of the BAR domain remains the same for both PH domains in an ACAP1BAR‑PH protein dimer. Lastly, the orientation of PH domain with respect to the BAR domain (α angles) in ACAP1BAR‑PH protein is defined as the angles between the orientation of the PH domain (blue arrows in Figure 1) and the three reference axes (yellow arrows in Figure 1) of the BAR domain (α1, α2, and α3). We first computed the α angles of the crystal structure of the ACAP1BAR‑PH protein dimer (PDB ID: 4NSW). Though the dimer is composed of two identical monomers, the α angles were found to be different for two PH domains. The results are as follows: α1, 21.92° and 5.58°; α2, 68.20° and 85.04°; α3, 87.76° and 87.46°. Theses angles implied that the PH domain may have flexible orientation angles relative to the BAR domain. The subsequent atomistic MD simulations would start from the crystal structures, therefore these angles also served as the initial α angles for the following dynamical evolution. We have carried out three atomistic MD simulations with a single ACAP1BAR‑PH protein dimer in ionic solvent (protein− solvent simulation, details in Methods). Flexibility of the PH domain in ACAP1BAR‑PH dimer was examined in the MD simulations. The temporal profiles of the α angles for each PH domain of the dimer for all the three simulations were shown in Figure 2. We note that all the α angles were disperse and indicate high dynamical flexibility of the loop linking the PH domain and the BAR domain. Although α1 angles started differently for the two PH domains in the crystal structure, they seemed to coincide in individual simulation ensembles (Table 1). However, the equilibrated α1 angles were different for three independent simulations (ranging from 11.62° to 22.58°), indicating that the dimer permits a variety of α1 angles. Similarly, the α2 angles also started from different angles for the two PH domains, then evolved to angles ranging from 71.82° to 96.00°. The α2 angles generally have larger uncertainties, reflected in larger standard deviations (SD), than those of the α1 angles, also indicating flexibility of the loop. Interesting, α3 angles of the two PH domains started from same initial angles, but diverged in some simulations and caused very different residual α3 angles. All the evidence show that the PH domains are loosely linked to the BAR domains. We note that transition between states with different protein structures might require at least tens of microseconds59 MD simulations. We have instead shown that ACAP1BAR‑PH protein dimer deviated from identical crystal structures and equilibrated to structures with very different orientation of the PH domain with respect to the BAR domain in multiple MD simulations. One of the PH Domains Rotates when Binding to the Membrane Asymmetrically. Then, we performed two independent atomistic MD simulations of one single ACAP1BAR‑PH protein dimer being placed above PIP2containing lipid bilayers (protein−lipid simulation, details in Methods Section). In both simulations, we have observed only one of the PH domains binding to the underneath membrane,

Figure 1. Definition of the orientation of PH domain with respect to the BAR domain. An ACAP1BAR‑PH protein dimer is shown with the BAR domain colored in blue and the PH domain in magenta. Three principle axes of the BAR domain are labeled shown as yellow arrows. Orientation of the PH domain corresponds to the principle axis of residue 345−361 (α5 helix) along the length of the helix and is shown as the blue arrow. The snapshot was rendered with VMD.49 3588

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Figure 2. Temporal profile of the orientation analysis of three protein−solvent simulations, namely simulation 1 (sim 1), 2 (sim 2), and 3 (sim 3). (A−C) α1, α2, and α3, angles. Profiles of the two PH domains of the ACAP1BAR‑PH protein dimer are shown separately (PH1 and PH2). Running averages were computed to show more clearly the evolution of the orientation angles.

Table 1. Orientation Analysis of the PH Domain in ACAP1BAR‑PH Protein Dimera α1

P-W

α2

P-M P-W

α3

P-M P-W

simulation 1

simulation 2

simulation 3

simulation 4

simulation 5

11.62 (4.32) 12.07 (6.69)

8.24 (4.43) 8.11 (4.29)

21.50 (4.81) 22.58 (5.59)

10.92 (6.36)

8.55 (3.59)

90.32 (6.66) 83.08 (8.26)

89.27 (5.87) 87.02 (6.44)

96.00 (5.99) 71.82 (5.35)

10.23 (4.30) 86.85 (9.06)

8.23 (3.76) 84.28 (5.35)

80.64 (4.53) 96.18 (5.81)

92.03 (6.93) 93.19 (4.82)

70.35 (4.57) 102.37 (3.79)

81.89 (5.78) 87.22 (7.59)

84.28 (4.70) 86.39 (3.38)

92.63 (4.09)

93.85 (3.44)

P-M

Simulations 1−3 were the protein-solvent simulations. Simulations 4−5 were the protein−membrane simulations. α1, α2, and α3 are the angles between the orientation of the PH domain (blue arrows in Figure 1) and the three reference axes (yellow arrows in Figure 1) of the BAR domain. PW refers to the PH domain interacting with solvent. P-M refers to the PH domain interacting with the membrane. Error estimation from standard deviation of the data is included in brackets. a

Figure 3. Temporal profile of the orientation analysis of two protein−lipid simulations, namely simulation 4 (sim 4) and 5 (sim 5). (A−C) α1, α2, and α3, angles. Profiles of the two PH domains of the ACAP1BAR‑PH protein dimer are shown separately as P-M (blue) and P-W (green). As one PH domain bound to the lipid bilayers, the another one remained unbound. Running averages were computed to show more clearly the evolution of the orientation angles.

to a much smaller range of angles compared to the case of protein in water. The α3 angles, in contrast, show very different equilibrated angles (in average of two PH domains, 93.24° for P-M and 86.81° for P-W), clearly distinguished the binding and nonbinding state of the PH domains (Table 1). Moreover, the P-M angles were seen to have smaller SD than those of the P-W angles in both protein−lipid and protein−solvent simulations. The smaller SD is intuitively caused by the interactions with the underneath membranes. As a result, the bound PH domain shows consistent orientations with small SD, while the unbound PH domain was associated with a different orientation with relatively larger SD of α3 angles. To conclude, introducing PIP2-containing lipid bilayers under ACAP1BAR‑PH protein dimer will cause asymmetrical

i.e., asymmetrical protein binding events. More importantly, with similar starting structures, different ends of the ACAP1BAR‑PH protein dimer were observed to bind to the membrane in the simulations, i.e., one end binds in simulation 1 and the another end binds in simulation 2. As the PH domains are loosely linked to the BAR domains, the PH domain is expected to explore a wide range of orientation angles. We note that the flexible loop between the BAR and PH domain may provide the chance for one of the PH domain to rotate to an angle facilitating the binding of protein to the lipid bilayers. We characterized the two ends into binding (P-M, P for protein, M for membrane) and nonbinding (P-W, W for water) ends. Figure 3 shows that, upon binding of the protein to the membrane, both α1 and α2 angles converged 3589

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Figure 4. Angle distribution of all the equilibrated data from Figures 2−3. (A−C) Protein−solvent simulations. (D−F) Protein−lipid simulations. Angles at the P-W state are shown in green, and P-M in blue. Bin sizes of 2° were used.

Figure 5. A simplified model of an ACAP1BAR‑PH protein dimer during membrane binding process. Snapshot of an ACAP1BAR‑PH dimer from (A) MD simulation of an ACAP1BAR‑PH dimer in ionic solvent. (B) MD simulation of an ACAP1BAR‑PH dimer on lipid bilayers. (A,B) Two monomers of the protein dimer are colored in a sense that PH domain of the green monomer interacting mostly with water and ion molecules (P-W state) and the blue one interacting with the membrane (P-M state). Similar to Figure 1, the blue arrows define the orientation of the PH domains; the three yellow arrows in (B), which were calculated from the principle axes of the BAR domains, define the orientation of the BAR domains. All snapshots were rendered with VMD.49

4D−F). The presence of lipids established bound states of the PH domain, therefore confined the dynamics of the bound protein dimer. In general, the fluctuations of the α angle in the bounded state decrease to 3−5° from 3 to 9° in unbounded states (Table 1). Especially for the α3 angle (Figure 4F), a deviation in P-M and P-W angles indicates strong interactions between the protein and nearby lipids, or even specific kind of lipids. By combining multiple all-atom MD simulations, orientation analysis has shown some molecular details of the membrane binding process of ACAP1BAR‑PH protein dimers. We note that as ACAP1 protein was previously reported to tubulate lipid liposomes. Well-structured lattices of ACAP1 protein upon binding to cell membrane were observed by cryoelectron microscopy (cryo-EM).32 Due to asymmetrical binding of ACAP1BAR‑PH protein dimer, formation of protein lattice would intrinsically introduce large area of curvatures, causing tubulation upon establishment of protein−lipid and protein− protein interactions. Recent studies on other BAR proteins60,61 has suggested factors, such as protein density, behind the complicated oligomerization events. For ACAP1 proteins, our orientation analysis might be able to provide valuable molecular information for the tubulation process. While this study focuses on study of protein−lipid interactions, details of protein− protein interactions require further studies in the future.

binding events, which associate with protein structural changes reflected by rotation of one of the PH domains with respect to the BAR domain. Orientation Analysis Revealed the Process of Protein−Membrane Binding. To understand protein binding to the membrane process from its configuration in the aqueous environment, we summarized the α angle distribution of all conducted MD simulations in Figure 4. We suggest that internal structural changes might have occurred when ACAP1BAR‑PH protein dimers interact with the lipids (Figure 5). First, when an ACAP1BAR‑PH protein dimer stands alone under physiological environment, both of the PH domains is at the unbound (P-W) state (Figure 5A). Without neighboring lipids, the PH domain of ACAP1BAR‑PH protein dimer was found to access a wide range of α angle (Figure 4A−C) with relatively large SD (up to 8° in Table 1). When placing PIP2containing membranes into the simulation system, ACAP1BAR‑PH protein dimer binds to the lipids asymmetrically and develops P-M and P-W states for the two PH domains (Figure 5B). Asymmetrical binding events have been observed in multiple independent protein−lipid simulations. It has been shown that both PH domains in an ACAP1BAR‑PH dimer are capable for binding to PIP2-containing lipid bilayers. Comparing the α angle to protein−solvent systems, protein−lipid systems certainly have more concentrated angle distribution due to the interactions between the protein and lipids (Figure 3590

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Figure 6. (A) Snapshot of a PH domain of the ACAP1BAR‑PH protein dimer from protein−lipid simulation 4. The PH domain is shown with cartoon representations in silver, except for the key residue loops. Three key residue loops defining the binding pocket interface were colored in lighter blue (Loop 1, residue 277−283), green (Loop 2, 300−305), and red (Loop 3, residue 321−327). Two bound PIP2 lipid head molecules are colored in orange (Pocket 1) and cyan (Pocket 2) to distinguish the binding pockets. Residues in contact with the PIP2 lipids are colored according to their side chain properties (magenta for negative, blue for positive, green for polar, and white for nonpolar). The BAR domain is shown with cartoon representations in transparent white. (B,C) Temporal profiles of separation between the pocket interface COM and the bound PIP2 lipid backbone phosphorus atoms. COM of Loop 1 and Loop 3 defines the Pocket 1 interface COM, and that of Loop 1 and Loop 2 defines the Pocket 2. (D) Number of contacts were counted every 50 ps for protein−lipid simulation 4 and then averaged over the number of frames. A contact is defined with a 12 Å cutoff between COM of the residue and the lipid backbone phosphorus atom. (E) The contact distribution in (D) projected onto the PH domain molecular surface, with blue corresponds to no contact and red corresponds to the most contacts. Loop 0, which is located between the two binding pockets, is circled. All snapshots were rendered with VMD.49

MD Simulations Reveal Two Binding Pockets of PH Domains. Analysis of orientation of the PH domains has pushed further our studies on the protein−lipid binding state of ACAP1BAR‑PH protein dimer. In agreement with the previously reported PIP2 specificity of ACAP1 protein,32 we have observed binding of PIP2 to PH domains in two protein−lipid simulations (Figure 6A). PIP2 binding events were facilitated by two binding pockets in the PH domain of ACAP1BAR‑PH protein dimer. In both simulations, we observed at least one PIP2 bound to each binding pocket. Two binding pockets are defined according to amino acid loops locate near where the PIP2 lipid bound. Three key residue loops were thus noted and colored in Figure 6A. Loop 1 (residue 277−283, β1/β2 loop), reported earlier by Pang et al.,32 inserts into the lipid bilayer and contributes most to the protein−lipid binding event. Loop 3 (residue 321−327, β5/β6 loop), together with Loop 1, composes the binding pocket 1 (Pocket 1) and interacts with a PIP2 lipid during the early stage of the simulations. Loop 2 (residue 300−305, β3/β4 loop), together with Loop 1, composes the binding pocket 2 (Pocket 2) and interacts with another PIP2 lipid after a period of time. Trajectories of the bound PIP2 were shown as the separation between the lipid backbone phosphate and the binding pockets (Figure 6B,C). Upon the binding of ACAP1BAR‑PH protein dimer to lipid bilayer underneath, Pocket 1 was found to establish stable

interactions between a nearby PIP2 lipid since a sharp decrease in pocket-lipid separation at ∼100 ns of simulation 1 and ∼20 ns of simulation 2 until the end of the simulation; while Pocket 2 gradually established such interactions throughout the simulation. Interestingly, although simulation 1 and 2 were independent simulations, the development of pocket−lipid interactions was very similar. Our orientation analysis suggested that the bound PH domain might undergo gradual rotation after inserting Loop 1 into lipid bilayers at ∼100 ns (Figure 2A−C). As a result, rotation of PH domain gradually decreased separation between the pocket and bilayer surface, facilitating binding of PIP2 lipid to Pocket 2. In order to identify key residues for the two pockets, Figure 6D show contacts between residues of the PH domain with nearby PIP2 lipids. Mapping of Figure 6D to molecular surface of a PH domain confirmed that the two binding pockets locate at two sides of the inserting Loop 1 (Figure 6E). Loop 1, 2, and 3 all have a number of contacts with PIP2 molecules. Specifically, residues D322, S323, E324, and R325 of Loop 3 have the highest amount of contacts among the loops and contribute mainly to Pocket 1. Residues N281 and T282 of Loop 1 and residues K299 and K300 of Loop 2 contribute mostly to the formation of Pocket 2. These results confirm the significance of electrostatic interactions between the PH domain of ACAP1 protein and PIP2 lipids. Identification of 3591

DOI: 10.1021/acs.jpcb.6b09563 J. Phys. Chem. B 2017, 121, 3586−3596

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The Journal of Physical Chemistry B

Figure 7. (A) Snapshot of Pocket 1 from protein−lipid simulation 4 at 255 ns. The blue sphere corresponds to the COM of Loop 1. The red sphere corresponds to the COM of two halves of Loop 3. The binding interface is represented as orange triangle with orange arrow indicating the normal. (B) Snapshot of Pocket 2 from protein−lipid simulation 4 at 418 ns. The blue sphere corresponds to the COM of Loop 1. The green sphere corresponds to the COM of two halves of Loop 2. The binding interface is represented as cyan triangle with cyan arrow indicating the normal. (A,B) Loop 1, 2, and 3 are colored in blue, green, and red, respectively. The rest residues are colored in white. Underneath lipid backbone phosphorus atoms are represented as transparent spheres. The black arrow indicates the normal of the membrane. (C,D) Orientation of Pocket 1 and Pocket 2 in protein−lipid simulation 4. (E,F) Orientation of Pocket 1 and Pocket 2 in protein−lipid simulation 5. (C−F) Running averages were computed to show more clearly the evolution of the angles. Color convention refers to the previous orientation analysis of PH domains in Figure 3.

key residues for the protein−lipid interactions not only hints for the binding interfaces, but also provides valuable information for future mutation experiments. Nature of the Two Binding Pockets of PH Domains. We examined the binding interface between the PH domains and membrane and found the PH domain rotates relative to the membrane normal direction during the PIP2 binding processes. Definition of the binding interface of the two binding pockets involves coarse-graining of the identified key residue loops. Loop 1 was coarse-grained into one bead according to the COM of the residues (Figure 7A,B). Loop 2 (in Figure 7A) and Loop 3 (in Figure 7B) were split into halves in order to define two beads for each loop. As a result, we were able to define a binding interface by connecting the three beads for each binding pocket (the orange triangle in Figure 7A and the blue triangle in Figure 7B). Dynamics of the binding interface could then be characterized by calculating angles between the normal

of the interface and the normal of the membrane surface (Figure 7C−F). In both simulations, Pocket 1 was seen to be heavily affected by the underneath membrane, reflected by the P-M angle having significantly smaller SD (3−5°) than the P-W angle (9−11°) (Figure 7C,E). In contrast, Pocket 2 underwent steady changes for both P-M and P-W angles, having relatively smaller SD (both 3−6°) (Figure 7D,F). This result supports our previous observation that PIP2 lipids follow a different but consistent trajectory heading the two binding pockets (Figure 6B−C). Interestingly, we observed that the orientation of Pocket 1 changed from ∼50° to 60° (Figure 7C) and that of Pocket 2 changed gradually from ∼80° to 60° (Figure 7D). This observation coincided with the fact that PIP2 lipids took time to approach Pocket 2 (Figure 6B) and the PH domain rotated to complete the binding process. Requirement in orientation of the PH domain has also been reported in various 3592

DOI: 10.1021/acs.jpcb.6b09563 J. Phys. Chem. B 2017, 121, 3586−3596

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Figure 8. (A) Snapshot of a PH domain at bound state with PIP2 lipids, from which we prepare all the initial structures for the US simulations. The atom coordinates were obtained from the last frame of protein−lipid simulation 4. The PH domain is shown in gray in cartoon representation; DOPC backbone phosphorus atoms of the lipid bilayers are represented by pink spheres. Two PIP2 lipids shown in licorice representation indicate the position of PIP2 lipids in two independent US setups. The orange PIP2 lipid represents the initial bound state of the Pocket 1/PIP2 setup; the cyan PIP2 lipid represents that of the Pocket 2/PIP2 setup. The black arrow indicates the direction of pulling during SMD simulations for preparing the subsequent US windows. The black dotted line indicates the separation used for restraints of US windows, which is the separation between protein COM and lower-layer lipid backbone phosphorus atoms COM. (B) PMF profiles for PH domain of ACAP1BAR‑PH protein dimer bound to PIP2 at Pocket 1 and Pocket 2. Protein−lipid separation was measured from the protein COM to the COM of lower layer lipid backbone phosphorus atoms along the membrane normal. Error estimates were obtained from bootstrap analysis.

Figure 9. Three-dimensional histograms of orientation angle of (A) the Pocket 1/PIP2 US (B) the Pocket 2/PIP2 US throughout the 15 US windows. The angle refers to the orientation of the two binding interfaces. The restrained separations are the window centers of US simulations which measured the separation between protein COM and lower-layer lipid backbone phosphorus atoms COM. The bin size is 1°.

studies10,12 in order to facilitate binding of lipids to different binding sites. Previous studies of GRP1 PH domain11,12 have revealed the same two binding pockets, in which a PH domain was placed alone and allowed to rotate freely, residues corresponding to Pocket 2 were reported to have the highest contact frequencies with bound PIP2 ligand, namely the “canonical site”. In order to find out whether the same difference preserves in PH domain of the ACAP1BAR‑PH protein dimer, we conducted multiple umbrella sampling simulations (Figure 8A) and generated PMF profiles of two pockets binding to PIP2 lipids, respectively (the blue and red profile in Figure 8B). Both profiles have a global minimum at a protein−lipid separation of ∼58 Å, with a well depth of −4.5 kcal mol−1 for Pocket 1 and −6.5 kcal mol−1 for Pocket 2. These profiles suggest favorable binding of both pockets to PIP2 with a preference for Pocket 2. Identification of two pockets and associated PIP2 preference is in agreement

with both previous experimental and computational results.9−12,62,63 In order to account for the specific binding to PIP2 lipids by PH domain, we have performed two more US simulations by replacing the PIP2 lipid at two binding pockets with POPS lipid, respectively. PMF profiles of two pockets binding to POPS lipids are also shown as yellow and purple plots in Figure 8B. The PMF of both pockets binding to POPS have a well depth of −2.0 kcal mol−1 and relative shorter range of coverage (about 6 Å), we demonstrate their preference for PIP2 over POPS lipid. Different well depths of the PMF profiles could be explained partly by the charged residues located inside the pockets. Pocket 1 consists of three positively charged residues: R275, K281, and R325; while Pocket 2 consists four positively charged residues: K274, R286, K299, and K336. Pocket 1 consists of one negatively charged residue E324, while Pocket 2 does not have any. Therefore, the less negative POPS lipids 3593

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process and binding interface orientations. More importantly, preference of the PH domain binding to PIP2 lipids over POPS lipids was also shown in the PMF results. Exploring the free energy surface of the protein−lipid systems revealed a much deeper well when a PIP2 lipid was in either binding pocket of the protein, compared to the case of POPS lipid.

(−1e) are expected to be electrostatically attracted by the charged residues less firmly, compared to PIP2 lipids (−3e). As a result, Pocket 2/PIP2 possessed the lowest well depth of PMF profile, Pocket 1/PIP2 came the second. Pocket 1/POPS and Pocket 2/POPS was seen to have similar and the shallowest PMF well. In addition to the PMF profile, our US simulations also provide further insights into nature of the binding pockets. Angle distributions of orientation of the binding interfaces during US simulations were given in Figure 9, in which the restrained separation covered a distance of ∼15 Å from the bound state. With a decrease in separations, Pocket 1 first spread over a wide range of orientation angles, then quickly converged to ∼60° when the PIP2 lipids came near the pocket (Figure 9A). Pocket 2 proved itself as a long-range interacting binding pocket, compared to Pocket 1, as the pocket orientation evolved gradually from ∼80° to ∼40° with decreasing restrained separations (Figure 9B). Difference in the effective range of the pockets also reflects in the PMF profiles, in which the Pocket 2/PIP2 well lifted over a longer distance of ∼2 Å than the Pocket 1/PIP2. We note that although both our energetic analysis and previous studies11,12 confirm that Pocket 2 is the preferred binding pocket for PIP2 lipids, Pocket 2 was found to bind PIP2 lipids later than Pocket 1 in both of our protein−lipid simulations (Figure 6B,C). It implies that multiple factors could lie behind protein−membrane binding events. As seen in Figure 6A, existence of the BAR domain in ACAP1BAR‑PH protein dimer might assist the binding, therefore confine the binding angle of the PH domain to cell membranes. As reported by Cui et al.,31 residues at the tip of the BAR domain might also involve in membrane binding. Preference binding by Pocket 2 then took effects by gradually rotating the PH domain and establishing interactions with a relatively far away PIP2 lipid. In this way, the ACAP1BAR‑PH protein dimers could quickly target and bind to desire regions of the cell membrane using Pocket 1, then reinforce such binding by strongly interacting with more PIP2 lipids using Pocket 2.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpcb.6b09563. Root-Mean Square Deviation (RMSD) profiles of the five MD simulations (namely sim1−5) in order to show the convergence of the simulations, convergence of the US simulations by calculating profiles of 30 ns intervals in 10 ns sliding blocks, histograms of the last 50 ns interval with PMF profiles (PDF)

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AUTHOR INFORMATION

Corresponding Author

*Tel: +(852) 3442 9978; E-mail: [email protected]. ORCID

Jun Fan: 0000-0001-8227-9671 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The authors thank the projects supported by the National Natural Science Foundation of China (21403182), the Research Grants Council of Hong Kong (CityU 21300014), and CityU grants (7004387 and 9680136). The authors would also like to acknowledge grants to F.S. by the Chinese Ministry of Science and Technology (2014CB910700) and the Strategic Priority Research Program of Chinese Academy of Sciences (XDB08030202). This research has used resources of the Oak Ridge Leadership Computing Facility at the Oak Ridge National Laboratory, which is supported by the Office of Science of the U.S. Department of Energy under Contract No. DE-AC05-00OR22725. This research has also used computing resources supported by Special Program for Applied Research on Super Computation of the NSFC-Guangdong Joint Fund (the second phase).

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CONCLUSION Our results provide insight into the structural dynamics of ACAP1BAR‑PH protein dimer when binding to cell membranes. Our simulations reveal that the PH domains of an ACAP1BAR‑PH protein dimer would undergo orientation changes during the membrane binding process. Such process was difficult to be caught in conventional experiments yet has suggested important residues for protein−lipid interactions. The list of dominating charged residues serve for the purpose of understanding and manipulating the membrane binding process by the PH domain of ACAP1BAR‑PH protein or other similar peripheral protein domains. A full understanding of membrane tubulation process will have to await explicit simulations of multiple ACAP1BAR‑PH protein dimers on PIP2containing membranes. However, the present simulations already reveal the principle protein structural deformation and key residue information upon protein−lipid interactions. In addition, protein−lipid interactions between ACAP1BAR‑PH protein dimer and PIP2-containing membranes have been studied by revealing two binding pockets on the PH domain. Preference for the two binding pockets is explained by PMF profiles generated by US simulations over separation of 12 Å between protein and lipids. Nature of the two binding pockets has been discussed in details with evolution of the binding

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REFERENCES

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