Scanning Tunneling Microscope Observation of the

Apr 26, 2015 - Taro Yamada,. ‡. Toshihide Kobayashi,. ‡ and Maki Kawai*. ,†. †. Department of Advanced Materials Science, The University of To...
0 downloads 0 Views 4MB Size
Download PDF
Article pubs.acs.org/Langmuir

Scanning Tunneling Microscope Observation of the Phosphatidylserine Domains in the Phosphatidylcholine Monolayer Soichiro Matsunaga,*,† Taro Yamada,‡ Toshihide Kobayashi,‡ and Maki Kawai*,† †

Department of Advanced Materials Science, The University of Tokyo, 5-1-5 Kashiwanoha, Kashiwa, Chiba 277-8561, Japan Lipid Biology Laboratory, RIKEN, 2-1 Hirosawa, Wako, Saitama 351-0198, Japan

‡

ABSTRACT: A mixed monolayer of 1,2-dihexanoyl-snglycero-3-phospho-L-serine (DHPS) and 1,2-dihexanoyl-snglycero-3-phosphocholine (DHPC) on an 1-octanethiolmodified gold substrate was visualized on the nanometer scale using in situ scanning tunneling microscopy (STM) in aqueous solution. DHPS clusters were evident as spotty domains. STM enabled us to distinguish DHPS molecules from DHPC molecules depending on their electronic structures. The signal of the DHPS domains was abolished by neutralization with Ca2+. The addition of the PS + Ca2+binding protein of annexin V to the Ca2+-treated monolayer gave a number of spots corresponding to a single annexin V molecule.

1. INTRODUCTION Phosphatidylserine (PS) is a major phospholipid in mammalian cells. In the plasma membrane, this lipid is mainly distributed on the inner cytoplasmic leaflet.1,2 It has been proposed that PS provides negatively charged lipid domains that work as a recruitment platform for specific proteins via ionic interaction. PS is exposed to the outer leaflet of the plasma membrane during apoptosis.1 In the inner leaflet monolayer, PS is in a phosphatidylethanolamine-rich environment, whereas in the outer leaflet PS is surrounded by a phosphatidylcholine (PC)rich membrane. The membrane distribution of PS in mammalian cells was recently examined by electron microscopy using a PS-specific protein of the GFP-tagged C2 domain of lactadherin.3 It is postulated that the lateral distribution pattern of lipids in membranes is determined primarily by physical lipid−lipid and lipid−protein interaction.4 Model membranes have been used to elucidate the molecular mechanisms underlying lateral lipid distribution.5−8 Lipids are visualized by using fluorescent lipid analogs or florescent indicators that reflect the physical properties of the membrane.8 Alternatively, label-free methods, such as light scattering, have also been employed.10,11 Scanning probe methods such as atomic force microscopy (AFM) and scanning tunneling microscopy (STM) are of particular interest because they visualize lipids on a nanometer scale under labelfree conditions. AFM primarily recognizes molecular size, and it can be utilized to visualize insulating materials.12,13 Thus, AFM is useful for distinguishing each lipid in a mixture of two or more lipids whose physical sizes are different.12,13 AFM has been extensively employed to image sphingolipid as higher domains in a supported phospholipid monolayer or bilayer in an aqueous environment.14−16 In contrast, a specific experimental setting is required to examine lipid mixtures of similarly sized lipids. STM detects the electronic state of molecules.17 © XXXX American Chemical Society

Thus, STM is expected to be able to distinguish two lipids that are similar in size but different in charge, such as PC and PS. Using STM, we have observed the fluidic motion and electrochemical phase transitions of a phospholipid monolayer spread on a hydrophobically modified substrate.18 Phospholipid nanoparticles have also been visualized by STM and have been shown to undergo a dynamic process of particle fusion initiated by a small antibiotic peptide.11 In this study, using in situ STM, we observed a mixed monolayer of 1,2-dihexanoyl-sn-glycero-3phosphocholine (DHPC) and 1,2-dihexanoyl-sn-glycero-3phospho-L-serine (DHPS) spread over an Au(111) singlecrystalline surface modified with 1-octanethiol and immersed in an aqueous NH4ClO4 buffer solution. The DHPS molecules were observed as spontaneously phase-separated domains embedded in a DHPC monolayer with an average diameter of approximately 6 nm. Neutralizing the charge of PS with Ca2+ abolished the PS signal. We also added annexin V to the system, which selectively binds to the PS + Ca2+ complex,19 and observed the protein binding to the lipid layer on the substrate at nanometer resolution.

2. MATERIALS AND METHODS 2.1. Materials. 1,2-Dihexanoyl-sn-glycero-3-phosphocholine (DHPC, purity >99%) and 1,2-dihexanoyl-sn-glycero-3-phospho-Lserine (DHPS > 99%) were products in the form of CHCl3 solutions from Avanti Polar Lipids Inc. CHCl3 (>99%), C2H5OH (spectroscopic grade), NH3 solution (ultrapure grade, 28.0−30.0%), HClO4 solution (ultrapure grade, 60.0−62.0%), CaCl2 (ultrapure grade, 95%), and 1octanethiol (1-C8H17SH) (>97%) were obtained from Kanto Chemicals, Japan. Full-length recombinant annexin V was obtained from Santa Cruz Biotechnology, Inc. (U.S.A., no. sc-4252). The gold Received: March 9, 2015 Revised: April 26, 2015

A

DOI: 10.1021/acs.langmuir.5b00859 Langmuir XXXX, XXX, XXX−XXX

Article

Langmuir crystal was prepared from an Au wire (diameter 1 mm, purity >99.999%, Furuya Metals, Japan). The electrodes for the STM electrochemical cell were made of Pt (wires of 1 and 0.25 mm diameter and a sheet of 0.1 mm thickness, >99.98%, Nilaco, Japan). 2.2. Methods. 2.2.1. Preparation of Samples. As the substrate for STM observation, we utilized a small (111) facet on a single-crystal gold bead by melting the Au wire in a flame.20,21 The facet was covered with a 1-octanthiol self-assembled monolayer by immersion in a 1 mM C2H5OH solution for 1 day.22−24 This hydrophobic surface was fixed on the in situ STM measurement cell. The desired phospholipidcontaining solutions for the formation of the phospholipid layer were poured into the cell for successive rounds of observation. On the basis of our experience, 18 the concentration of phospholipids in the solution needs to be approximately 2 mM to form layers detectable by STM. The critical micelle concentration of DHPC is 15 mM25 and that of DHPS should be higher than 15 mM,25 and thus DHPC and DHPS are dispersed in the solution as isolated molecules, with no aggregation anticipated in the 2 mM solution. All of the phospholipids were stored in CHCl3 at −30 °C. A portion of the CHCl3 solution was taken up in the desired volume into a small glass vial, and CHCl3 was allowed to evaporate under N2 gas flow. After the evaporation process, a calculated amount of buffer solution was poured into this vial. The solution was vortex mixed and sonicated in an ultrasonic bath (US102, SND, Japan) for 10 min. This solution was poured into the STM cell in which the hydrophobic 1-octanthiol/ Au(111) had been preset. The phospholipid molecules adsorbed on the substrate by the hydrophobic interaction and formed a monolayer on the surface. 2.2.2. In Situ Scanning Tunneling Microscope (in Situ STM). All of the in situ STM images were obtained using a Nanoscope E (Veeco Instruments Inc.). We utilized a commercial Pt/Ir (80:20) tip (Veeco). The tip was coated with nail polish to minimize the Faradaic current in the electrolyte. The measurement was performed in a Teflon-PFA (PerFluoroAlkoxyethylene) cell (liquid content volume = 2 mL), assembled with a Pt plate (6 mm × 20 mm) as the counter electrode and a Pt wire as the reference electrode. An NH4ClO4 buffer (0.05 M, pH 7.0 ± 0.1, prepared by neutralization of NH3 and HClO4 aqueous solutions) was used as the supporting electrolyte. The Pt quasireference electrode was electrochemically cleaned and calibrated with an Ag/AgCl electrode in the same solution. The substrate potential was always potentiostatically controlled at 0.22 V vs Ag/AgCl, which is the open circuit potential for the working electrode, that is, the model cell membrane on Au(111) in the lipid-containing solution. This is to avoid unexpected electrochemical reactions. In the STM images shown, the brighter area indicates that the STM tip is far from the sample surface. Because the images are taken in constant-current mode, the distance between the tip and the sample depends on the conductivity of the tunneling electron between the tip and the sample; therefore, STM is able to detect not only the physical height of sample objects but also their electronic structure.17 In this article, we use the word “height” as the distance between the tip and the sample. To interpret the experimental results, a density-function-theory (DFT) calculation was performed with the aid of a commercial program package26−28 (Accelrys DMol3 Ver. 4.0) installed in the RIKEN Integrated Cluster of Clusters (RICC) supercomputing system. The geometric structure was energetically optimized with a DFT molecular orbital calculation with the double-numerical plus dfunction basis set (DND option) for all electrons. The local density approximation (LDA) Perdue−Wang (PWC) functional was employed. The orbital wave function was visualized for each of the energy eigenvalues on the basis of the optimized molecular structure.

Figure 1. In situ STM images of the phospholipid monolayer on the substrate. The scanned area is 100 × 100 nm2. The left image was of a DHPC + DHPS mixed monolayer, and the concentration of phospholipids was DHPC/DHPS = 1.5 mM/0.5 mM. The right image is of a pure DHPC monolayer on the substrate, and the concentration of DHPC was 1.5 mM. The sample electrode potential was adjusted to 0 V vs Pt, and the tip potential was −0.4 V vs Pt. The tunneling current was 1.0 nA.

for DHPS and 1.5 mM for DHPC. A number of highconductivity spots were evident on flat terraces. The average size and height of the spots were 6.1 ± 1.0 and 0.4 ± 0.1 nm, as measured from the terrace level, respectively. The image on right in Figure 1 is an in situ STM image of a pure DHPC monolayer on the substrate. Similar to our previous report,18 a flat layer was formed on the substrate. Holes in the layer are composed of a defective area in the DHPC layer and vacancy islands (VIs) on the thiol-modified gold substrate. Characterization of the images of the DHPC monolayer was provided in our previous report.18 As shown in Figure 1, the spots were observed only in the DHPS-containing system. In contrast, the defective area in the phospholipid layer and VIs was detected in both images, indicating that, similar to the pure DHPC monolayer, the DHPS and DHPC molecules formed a mixed monolayer on the substrate. To estimate the composition of these spots, we changed the ratio of DHPS and DHPC in the solution and mixed monolayers. Figure 2 shows the in situ STM images of these monolayers. The concentrations of phospholipids in the solution were (A) DHPS/DHPC = 0.1 mM/2.0 mM, (B)

Figure 2. In situ STM images of the DHPS + DHPC monolayer on the substrate. The scanned area is 50 × 50 nm2. The concentrations of DHPS and DHPC in each image are shown in the table below. The sample electrode potential was adjusted at 0 V vs Pt, and the tip potential was −0.4 V vs Pt. The tunneling current was 1.0 nA. The graph below shows the relationship between the DHPS/(DHPS + DHPC) in the solution and the bright area ratio observed by in situ STM images.

3. RESULTS 3.1. Observation of the DHPS + DHPC Mixed Layer Using in Situ STM. In situ STM images obtained after the formation of the DHPS + DHPC monolayer on the 1octanethiol-modified-Au(111) substrate are shown in Figure 1. The phospholipid concentration in the solutions was 0.5 mM B

DOI: 10.1021/acs.langmuir.5b00859 Langmuir XXXX, XXX, XXX−XXX

Article

Langmuir 0.15 mM/1.5 mM, and (C) 0.5 mM/1.5 mM. As the ratio of DHPS in the solution increased, the area occupancy of the spots also increased. The table and graph in Figure 2 show the relationship between the ratio of DHPS in the solution and the brighter area in the in situ STM images. To measure the area, we used more than four in situ STM images (200 × 200 nm2) in each case, with the average values calculated by measuring more than 50 spots. Remarkably, the fraction of the spots over the total lipid monolayer area was proportional to the molar fraction of DHPS in the solution within the range of the DHPS concentrations tested. It is thus highly likely that the spots were composed of DHPS-rich lipid domains. To examine whether the spot is an aggregation of lipid molecules or a result of the quick fluctuation of a single lipid molecule, we measured the same area with the scanning speed changing from 0.15 nm/ms to 1.5 nm/ms. The observed sizes and features of the spots were not affected by the change in scanning speed. The result suggests that the observed spot size corresponds to the size of the DHPS-rich domain in the monolayer. The sizes, features, and position of the DHPS-rich domains did not drastically change by repeated STM scans at intervals of a few minutes. These results indicate that stable DHPS-rich domains formed in the mixed monolayer. 3.2. Ca2+ Effect on the DHPS + DHPC Mixed Layer. PS strongly binds with Ca2+.29 We examined the effect of Ca2+ on the STM images of the DHPS + DHPC mixed monolayer. We prepared three sets of DHPS + DHPC monolayers (the initial concentrations of DHPS is 0.15 mM, and that of DHPC is 1.5 mM). The in situ STM images of the DHPS + DHPC layers for each time are shown in Figure 3, row A. Then, we added the Ca 2+ solution to the STM measurement cell at Ca 2+ concentrations of 0.05, 0.075, and 0.4 mM, respectively. Row B of Figure 3 shows the in situ STM images immediately after the addition of Ca2+. The images were acquired from the same sample but over a different area. The images in row C of Figure 3 were acquired after several STM scans to facilitate the mixing of Ca2+. We made consecutive scans between B and C for each Ca2+ concentration, and images C-1, C-2, and C-3 were obtained from almost the same area as for B-1, B-2, and B-3, respectively. Adding Ca2+ at 0.05 mM did not notably change the STM image as shown in Figure 3, column 1. At 0.075 and 0.4 mM (columns 2 and 3), the bright spots eventually disappeared. The flat features of the phospholipid layer did not in general change drastically except for the bright spots being invisible. This concentration-dependent effect of Ca2+ also supports the present interpretation that the spots are DHPS-rich domains. Because STM detects the electronic states of the molecule, it is likely that the binding of Ca2+ neutralizes the negative charge of DHPS, so STM was unable to distinguish DHPS + Ca2+ and DHPC. However, considering the high water solubility of short-chain phospholipids, we cannot exclude the possibility that Ca2+ extract DHPS molecules from the monolayer. 3.3. Annexin V Adsorption on the Layer: The Presence of DHPS in the Homogeneous Layer. Annexin V binds the PS + Ca2+ complex.30−33 To examine whether DHPS remained after the addition of Ca2+, we added annexin V to the Ca2+treated DHPS + DHPC monolayer. Figure 4A shows a mixed phospholipid monolayer (DHPC/DHPS = 2.0 mM/0.1 mM, the same composition as in Figure 2C). Approximately 6-nmdiameter DHPS-rich domains were observed as spots. Figure 4B was recorded after the addition of 0.4 mM Ca2+. The

Figure 3. In situ STM images of the DHPC + DHPS monolayer after the addition of Ca2+. The scanned area is 50 × 50 nm2. Three sequences of experiments were demonstrated. 1, 2, and 3 show the number of sequences. In a sequence for each experiment, we observed the same sample. A-1, A-2, and A-3 are the images without Ca2+. The concentrations of DHPC and DHPS were 1.5 and 0.15 mM. B-1, B-2, and B-3 are the images after the addition of Ca2+. The concentrations of added Ca2+ was 0.05 mM in B-1, 0.075 mM in B-2, and 0.4 mM in B-3. Panels A and B were not obtained from the same area. C-1, C-2, and C-3 were obtained after several STM scans at each concentration of Ca2+; therefore, panels B and C in the same sequence were obtained from almost the same area.

Figure 4. In situ STM images of the DHPC + DHPS monolayer in the presence of Ca2+ and annexin V. The scanned area is 50 × 50 nm2. The concentrations of DHPC and DHPS are 2.0 and 0.1 mM, respectively. (A) DHPC and DHPS mixed layer, (B) obtained after the addition of Ca2+ to the DHPC + DHPS monolayer, and (C) obtained after the addition of 1 μg/mL Annexin V to DHPC + DHPS + Ca2+. Bright areas in (C) remained even after many scans over the same area. C

DOI: 10.1021/acs.langmuir.5b00859 Langmuir XXXX, XXX, XXX−XXX

Article

Langmuir domains disappeared as described in section 3.3. Then 1 μg/ mL of annexin V was added to the buffer solution. Figure 4C was immediately obtained after the addition of annexin V. As shown in Figure 4C, the spots were observed again. These spots did not disappear after any further addition of Ca2+. The average size of the spots was 5.0 ± 0.4 nm, and the average height was 0.25 ± 0.03 nm. Figure 5 shows the in situ STM images with the different ratio of DHPS and DHPC after the successive addition of Ca2+

Figure 6. In situ STM images of DHPC monolayers in the presence of Ca2+ and annexin V. This is the blank test of the annexin V effect on PS and Ca2+. The scanned area is 100 × 100 nm2. The concentration of DHPC was 1.5 mM. (A) DHPC monolayer on the substrate, (B) obtained after the addition of Ca2+ to (A), and (C) obtained after the addition of 1 μg/mL annexin V to (B).

from PC by molecular size. Indeed, in a previous AFM study by Vie et al.,35 a mixed lipid layer consisting of similarly molecularsized DOPC and DOPS, which have the same fatty acid composition, was observed as a homogeneous layer, and DOPS was not distinguishable. In the present STM experiment, the DHPS-rich domains were observed as higher-contrast spots than the background DHPC monolayer. Although DHPS and DHPC are similar to each other in terms of molecular size and structure, we actually observed a distinctive STM contrast of DHPS domains against the DHPC monolayer. The contrast observed in STM images is of course strongly related to the mechanism of tunnelingcurrent imaging. Our observation was performed in aqueous solution, in which the path of imaging electrons must deviate from that in the simple vacuum environment. The path may be serially divided into the tip−sample gap and the organic layer over the conductive substrate. The current through the tip−sample gap in our aqueous environment is the quantum mechanical tunneling current, similar to the case in vacuum. This is clear when we see the abundant examples of angstrom-scale STM images of organic molecules recorded in vacuum, in air, and in aqueous solution under essentially the same electric conditions. In most of those cases, the STM tip-to-sample bias voltage is set at a few hundred mV, and the tip current is less than a few nA. The Faradaic current in the electrolytic solution is excluded as the imaging current in the present case. Our STM observations were performed with a small tip-to-substrate voltage within the electric double-layer region. It is unlikely that the ion current is the main current for the imaging.36−39 The detectably high electronic current has not yet been completely understood for biological molecules on metal surfaces immersed in aqueous solution. Although even large molecules are conductive enough for in situ STM in aqueous solution, the same molecules are highly resistive in vacuum, in which STM imaging is hindered. This topic is often addressed in the literature, such as in refs 36 and 37. In short, biological molecules are in general heavily hydrated in electrolytic solution, in which many kinds of ionic and polar, neutral species are densely gathering. The electric current through a layer composed of such a mixture should be considered in such a context. The contrast of the DHPS domains in the DHPC background originates from the electronic state of individual DHPS and DHPC molecules. We made a simple consideration according to our own molecular orbital calculation result using commercial package Dmol3. Our simulation exhibited the

Figure 5. In situ STM images of the DHPC + DHPS monolayer in the presence of Ca2+ and annexin V. The scanned area is 100 × 100 nm2. The concentration of DHPC and DHPS were (A) DHPC/DHPS = 2.0 mM/0.1 mM and (B) DHPC/DHPS = 2.0 mM/0.2 mM, respectively. The concentrations of Ca2+ and annexin V were 0.4 mM and 1 μg/mL in both A and B.

and annexin V. The concentrations of DHPC, Ca2+, and annexin V were fixed, and the concentration of DHPS was twice as high in Figure 5B as in Figure 5A. Clearly, there are more spots in Figure 5B than in Figure 5A. The average diameter of the spots, 5 nm, matches the diameter of a single annexin V molecule,34 suggesting that the spot corresponds to a single protein that is bound to the complex of DHPS and Ca2+ within the monolayer. The desorption of annexin V from the layer was not observed even after repeated STM scans. We quantified the average occupancy of the annexin V area in the STM images represented in Figure 5 using more than four 50 × 50 nm2 frames. In Figure 5A (DHPC/DHPS = 2.0 mM/0.1 mM), the ratio of the bright area/whole frame was 0.04 ± 0.02 (average ± standard deviation), and in Figure 5B (DHPC/DHPS = 2.0 mM/0.2 mM), it was 0.11 ± 0.06. The area occupancy approximately matches the DHPS part in DHPS + DHPC. This result suggests that the DHPS-rich domains were bound to annexin V molecules. These results also indicate that DHPS existed in the monolayer during the course of Ca2+ and annexin V treatment. To confirm the selective binding of annexin V to the DHPS + Ca2+ complex, we examined the effect of Ca2+ and annexin V on the DHPC monolayer. Figure 6A shows an in situ STM image of the DHPC monolayer, Figure 6B is an image after the addition of Ca2+, and Figure 6C is an image after the further addition of annexin V. The concentrations of components were the same as those in Figure 5. We did not observe any change in the DHPC layer after the addition Ca2+ and annexin V.

4. DISCUSSION In the present study, we showed that STM is capable of distinguishing the DHPS molecules from the DHPC monolayer background. The DHPS and DHPC molecules have almost the same size. Both have two six-carbon fatty acids and a glycerophosphate group. The only difference is the choline and serine groups. Hence, it might be difficult to distinguish PS D

DOI: 10.1021/acs.langmuir.5b00859 Langmuir XXXX, XXX, XXX−XXX

Article

Langmuir

intrinsic factors, such as molecular size and electron conductivity, extrinsic factors, such as the molecular diffusion rate and STM scanning rate, also affect the detection of molecules by STM. In our simulation, various clustering patterns of two DHPS anions and Ca2+ can be postulated and optimized as energetic local minima by DFT molecular orbital calculations. Such structures exhibit LUMO wrapping around the glycerol/ carbonyl parts of the two DHPS molecules and the HOMO of the molecules located around the serine part. The spatial distributions of these orbitals are similar to those of DHPC. This might make a DHPS −2·Ca 2+ cluster and DHPC indistinguishable by STM observation.

lowest unoccupied molecular orbital (LUMO) localized around the glycerol/carbonyl part of DHPC and the highest occupied molecular orbital (HOMO) of the DHPC located around the choline part, whereas the LUMO of a single DHPS−·NH4+ cluster wrapped around the serine headgroup and the HOMO of the cluster localized around the glycerol/carbonyl parts. This crucial difference in the molecular orbitals may be reflected in the STM contrast, although the calculation was performed for a single molecule/cluster placed in a vacuum despite the much more complex reality of condensed phospholipid layers on the modified Au surface immersed in an electrolytic solution. In the DHPS + DHPC mixed monolayer, the DHPS molecules spontaneously formed 6 nm clusters. A molecular dynamics simulation estimated that this cluster contains roughly 50 PS molecules.40 In previous cases of macroscopic observation using spin resonance and calorimetry,41−43 mixtures of PS and PC were initially composed into homogeneous fluids, and then the addition of Ca2+ induced a phase transition from a fluidic phase to a gel phase. In addition, a Ca2+ induced micrometer-scale PS domain was visualized in the DPPC + DPPS Langmuir−Blodgett layer.44 On the other hand, the formation of tiny PS clusters even without Ca2+ has been predicted in computer simulation.45 In this study, we showed that DHPS molecules form nanometer-scale clusters. In previous reports the molecular-scale cluster formation has been explained by an excess free energy of the nonideal mixing of PS and PC.45 The formation of a PS-rich cluster can also be explained from the viewpoint of the lateral diffusion coefficient of the phospholipid layer spreading on the substrate. In our previous observation of the DHPC/1-octanethiol/Au(111) in aqueous media,18 the diffusion coefficient of the DHPC monolayer was ∼10−15 m2/s, and this value was 3 orders of magnitude smaller than that of the bilayer at ∼10−12 m2/s.46 The reduced fluidity of the phospholipid monolayer on alkanethiol-modified Au(111) was also studied using molecular dynamics simulation.47,48 One explanation suggested for the reduction in fluidity is that one leaflet of the bilayer consists of highly ordered and closely packed alkyl chains.18 When the fluidity is reduced, the lateral motion of phospholipids is suppressed and the phospholipid molecules tend to form a gellike phase. Therefore, both DHPS and DHPC molecules exhibit a gel-like phase on the substrate. The gel−liquidcrystalline phase-transition temperature of PS is higher than that of PC with the same fatty acid composition,25 and thus the PS−PS interaction is stronger than that the PC−PC interaction. This leads to the results that PS molecules aggregate more easily than PC molecules, and the formation of DHPS clusters occurs in DHPC even though the fatty acids are very short. The addition of Ca2+ to the DHPS + DHPC monolayer abolished the DHPS signal. The disappearance of the DHPS signal took place when the Ca2+ concentration exceeded 0.075 mM, which was half of the DHPS concentration. The result suggests that divalent Ca2+ is neutralized by two negatively charged monovalent DHPS molecules. The binding of annexin V after Ca2+ treatment of the monolayer excluded the possibility that it was the Ca2+ treatment that extracted DHPS from the monolayer. The domain disappearance was enhanced by STM scanning, suggesting that STM scanning accelerated the mixing of the solution and the formation of complexes of Ca2+ and DHPS−. At higher Ca2+ concentrations, the DHPS-rich domains disappeared immediately after the addition of Ca2+. These results indicate that in addition to

5. CONCLUSIONS In this report, in situ STM was performed to observe a mixed monolayer of negatively charged DHPS and neutral DHPC. We visualized DHPS-rich domains in the mixed monolayer, the disappearance of these domains by the addition of Ca2+, and annexin V binding to DHPS and Ca2+ complexes in the monolayer. As a result of these nanometer-scale visualizations, we report three new findings as described below. 1. STM enabled us to recognize the DHPS molecules from the neutral DHPC background layer. DHPS-rich domains that were 6 nm in diameter formed in the fluidic DHPC + DHPC mixed monolayer on the substrate. 2. The DHPS-rich domains disappeared in the presence of Ca2+. This can be explained by the neutralization of DHPS by Ca2+ binding. 3. The presence of DHPS molecules in the monolayer after the addition of Ca2+ was confirmed by annexin V binding to the layer. The binding of single annexin V molecules to the DHPS-rich domains was visualized. In this work, it was demonstrated that in situ STM is a powerful technique for distinguishing similarly sized phospholipids depending on their electronic structures. This is one of the advantages of STM compared to AFM in terms of the observation of biological objects. Nanometer-scale spatial resolution and label-free observation are characteristics of in situ STM which are applicable to studies of various lipid−lipid and lipid−protein interactions.

■

AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected] (S.M). *E-mail: [email protected] (M.K.). Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS This study was financially supported in part by the RIKEN President Discretionary Fund (2004−2006, 2010−2012), the Lipid Dynamics Program and Integrated Lipidology Program of RIKEN, and Grants-in-Aid for Scientific Research on the Promotion of Novel Interdisciplinary Fields Based on Nanotechnology and Materials from the Ministry of Education, Culture, Sports, Science and Technology of Japan. This work was also supported by KAKENHI (grant nos. 19360024 and 25293015). S.M. acknowledges support from Research Fellowships for Young Scientists of the Japan Society for the Promotion of Science. We thank RIKEN, Japan, for an E

DOI: 10.1021/acs.langmuir.5b00859 Langmuir XXXX, XXX, XXX−XXX

Article

Langmuir

(22) Twardowski, M.; Nuzzo, R. G. Molecular recognition at model organic interfaces: Electrochemical discrimination using self-assembled monolayers (SAMs) modified via the fusion of phospholipid vesicles. Langmuir 2003, 19, 9781−9791. (23) Twardowski, M.; Nuzzo, R. G. Phase dependent electrochemical properties of polar self-assembled monolayers (SAMs) modified via the fusion of mixed phospholipid vesicles. Langmuir 2004, 20, 175− 180. (24) Doyle, A. W.; Fick, J.; Himmelhaus, M.; Eck, W.; Graziani, I.; Prudovsky, I.; Grunze, M.; Maciag, T.; Neivandt, D. J. Protein deformation of lipid hybrid bilayer membranes studied by sum frequency generation vibrational spectroscopy. Langmuir 2004, 20, 8961−8965. (25) http://avantilipids.com/. (26) Delley, B. An all-electron numerical method for solving the local density functional for polyatomic molecules. J. Chem. Phys. 1990, 92, 508−517. (27) Delley, B. Fast calculation of electrostatics in crystals and large molecules. J. Phys. Chem. 1996, 100, 6107−6110. (28) Delley, B. From molecules to solids with the DMol3 approach. J. Chem. Phys. 2000, 113, 7756−7764. (29) Newton, C.; Pangborn, W.; Nir, S.; Papahadjopoulos, D. Specificity of Ca2+ and Mg2+ binding to phosphatidylserine vesicles and resultant phase changes of bilayer membrane structure. Biochim. Biophys. Acta, Biomembr. 1978, 506, 281−287. (30) Gerke, V.; Creutz, C. E.; Moss, S. E. Annexins: Linking Ca2+ signalling to membrane dynamics. Nat. Rev. Mol. Cell Biol. 2005, 6, 449−461. (31) van Genderen, H. O.; Kenis, H.; Hofstra, L.; Narula, J.; Reutelingsperger, C. P. M. Extracellular annexin A5: Functions of phosphatidylserine-binding and two-dimensional crystallization. Biochim. Biophys. Acta, Mol. Cell Res. 2008, 1783, 953−963. (32) Wang, J. L.; Gray, R. M.; Haudek, K. C.; Patterson, R. J. Nucleocytoplasmic lectins. Biochim. Biophys. Acta, Gen. Subj. 2004, 1673, 75−93. (33) Demange, P.; Voges, D.; Benz, J.; Liemann, S.; Gottig, P.; Berendes, R.; Burger, A.; Huber, R. annexin-V − the key to understanding ion selectivity and voltage regulation. Trends Biochem. Sci. 1994, 19, 272−276. (34) http://www.rcsb.org/pdb/home/home.do. (35) Vie, V.; Legardinier, S.; Chieze, L.; Le Bihan, O.; Qin, Y.; Sarkis, J.; Hubert, J. F.; Renault, A.; Desbat, B.; Le Rumeur, E. Specific anchoring modes of two distinct dystrophin rod sub-domains interacting in phospholipid Langmuir films studied by atomic force microscopy and PM-IRRAS. Biochim. Biophys. Acta, Biomembr. 2010, 1798, 1503−1511. (36) Zhang, J.; Kuznetsov, A. M.; Medvedev, I. G.; Chi, Q.; Albrecht, T.; Jensen, P. S.; Ulstrup, J. Single-molecule electron transfer in electrochemical environments. Chem. Rev. 2008, 108, 2737−2791. (37) Zhang, J.; Chi, Q.; Hansen, A. G.; Jensen, P. S.; Salvatore, P.; Ulstrup, J. Interfacial electrochemical electron transfer in biology towards the level of the single molecule. FEBS Lett. 2012, 586, 526− 535. (38) Guckenberger, R.; Heim, M.; Cevc, G.; Knapp, H. F.; Wiegrabe, W.; Hillebrand, A. Scanning tunneling microscopy of insulators and biological specimens based on lateral conductivity of ultrathin water films. Science 1994, 266, 1538−1540. (39) Fan, F.-R.; Bard, A. J.; Guckenberger, R.; Heim, M. STM on wet insulators: electrochemistry or tunneling? Science 1995, 270, 1849− 1852. (40) Pandit, S. A.; Berkowitz, M. L. Molecular Dynamics Simulation of Dipalmitoylphosphatidylserine Bilayer with Na+ Counterions. Biophys. J. 2002, 82, 1818−1827. (41) Ohnishi, S.; Ito, T. Calcium-induced phase separations in phosphatidylserine-phosphatidylcholine membranes. Biochemistry 1974, 13, 881−887. (42) Silvius, J. R.; Gagne, J. Calcium-induced fusion and lateral phase separations in phosphatidylcholine-phosphatidylserine vesicles −

allocation of supercomputing resources on the RIKEN Integrated Cluster of Clusters (RICC) system.

■

REFERENCES

(1) Kay, J. G.; Grinstein, S. Phosphatidylserine-mediated cellular signaling. In Lipid-Mediated Protein Signaling; Springer: 2013; pp 177− 193. (2) Hullin-Matsuda, F.; Taguchi, T.; Greimel, P.; Kobayashi, T. In Lipid compartmentalization in the endosome system, Semin. Cell Dev. Biol. 2014, 48−56. (3) Fairn, G. D.; Schieber, N. L.; Ariotti, N.; Murphy, S.; Kuerschner, L.; Webb, R. I.; Grinstein, S.; Parton, R. G. High-resolution mapping reveals topologically distinct cellular pools of phosphatidylserine. J. Cell Biol. 2011, 194, 257−275. (4) Lingwood, D.; Simons, K. Lipid rafts as a membrane-organizing principle. Science 2010, 327, 46−50. (5) Edidin, M. The state of lipid rafts: from model membranes to cells. Annu. Rev. Biophys. Biomol. Struct. 2003, 32, 257−283. (6) London, E. How principles of domain formation in model membranes may explain ambiguities concerning lipid raft formation in cells. Biochim. Biophys. Acta, Mol. Cell Res. 2005, 1746, 203−220. (7) Sonnino, S.; Prinetti, A. Membrane domains and the “lipid raft” concept. Curr. Med. Chem. 2013, 20, 4−21. (8) Ramstedt, B.; Slotte, J. P. Sphingolipids and the formation of sterol-enriched ordered membrane domains. Biochim. Biophys. Acta, Biomembr. 2006, 1758, 1945−1956. (9) Veatch, S. L.; Keller, S. L. Seeing spots: complex phase behavior in simple membranes. Biochim. Biophys. Acta, Mol. Cell Res. 2005, 1746, 172−185. (10) Wydro, P. Sphingomyelin/phosphatidylcholine/cholesterol monolayers-analysis of the interactions in model membranes and Brewster Angle Microscopy experiments. Colloids Surf., B 2012, 93, 174−179. (11) Matsunaga, S.; Matsunaga, T.; Iwamoto, K.; Yamada, T.; Shibayama, M.; Kawai, M.; Kobayashi, T. Visualization of Phospholipid Particle Fusion Induced by Duramycin. Langmuir 2009, 25, 8200−8207. (12) Goksu, E. I.; Vanegas, J. M.; Blanchette, C. D.; Lin, W.-C.; Longo, M. L. AFM for structure and dynamics of biomembranes. Biochim. Biophys. Acta, Biomembr. 2009, 1788, 254−266. (13) García-Sáez, A. J.; Schwille, P. Surface analysis of membrane dynamics. Biochim. Biophys. Acta, Biomembr. 2010, 1798, 766−776. (14) Giocondi, M.-C.; Boichot, S.; Plenat, T.; Le Grimellec, C. Structural diversity of sphingomyelin microdomains. Ultramicroscopy 2004, 100, 135−143. (15) Wang, T.; Shogomori, H.; Hara, M.; Yamada, T.; Kobayashi, T. Nanomechanical recognition of sphingomyelin-rich membrane domains by atomic force microscopy. Biochemistry. 2011, 51, 74−82. (16) Ros, U.; Edwards, M. A.; Epand, R. F.; Lanio, M. E.; Schreier, S.; Yip, C. M.; Alvarez, C.; Epand, R. M. The sticholysin family of poreforming toxins induces the mixing of lipids in membrane domains. Biochim. Biophys. Acta, Biomembr. 2013, 1828, 2757−2762. (17) Wiesendanger, R. Scanning Probe Microscopy and Spectroscopy: Methods and Applications; Cambridge University Press: Cambridge, 1994. (18) Matsunaga, S.; Yokomori, R.; Ino, D.; Yamada, T.; Kawai, M.; Kobayshi, T. EC-STM observation on electrochemical response of fluidic phospholipid monolayer on Au(111) modified with 1octanethiol. Electrochem. Commun. 2007, 9, 645−650. (19) Walker, J.; Boustead, C.; Koster, J.; Bewley, M.; Waller, D.; Annexin, V. a calcium-dependent phospholipid-binding protein. Biochem. Soc. Trans. 1992, 20, 828−833. (20) Hamelin, A.; Morin, S.; Richer, J.; Lipkowski, J. Adsorption of pyridine on the (110) face of silver. J. Electroanal. Chem. 1989, 272, 241−252. (21) Hamelin, A.; Morin, S.; Richer, J.; Lipkowski, J. Adsorption of pyridine on the (311) face of silver. J. Electroanal. Chem. 1990, 285, 249−262. F

DOI: 10.1021/acs.langmuir.5b00859 Langmuir XXXX, XXX, XXX−XXX

Article

Langmuir correlation by calorimetric and fusion measurements. Biochemistry 1984, 23, 3241−3247. (43) Ito, T.; Ohnishi, S. I. Ca2±induced lateral phase separations in phosphatic acid-phosphatidylcholine membranes. Biochim. Biophys. Acta 1974, 352, 29−37. (44) Ross, M.; Steinem, C.; Galla, H. J.; Janshoff, A. Visualization of chemical and physical properties of calcium-induced domains in DPPC/DPPS Langmuir-Blodgett layers. Langmuir 2001, 17, 2437− 2445. (45) Huang, J.; Swanson, J. E.; Dibble, A.; Hinderliter, A. K.; Feigenson, G. W. Nonideal mixing of phosphatidylserine and phosphatidylcholine in the fluid lamellar phase. Biophys. J. 1993, 64, 413−425. (46) Tero, R.; Watanabe, H.; Urisu, T. Supported phospholipid bilayer formation on hydrophilicity-controlled silicon dioxide surfaces. Phys. Chem. Chem. Phys. 2006, 8, 3885−3894. (47) Tu, K.; Tobias, D. J.; Klein, M. L. Constant pressure and temperature molecular dynamics simulation of a fully hydrated liquid crystal phase dipalmitoylphosphatidylcholine bilayer. Biophys. J. 1995, 69, 2558−2562. (48) Tarek, M.; Tu, K.; Klein, M. L.; Tobias, D. J. Molecular dynamics simulations of supported phospholipid/alkanethiol bilayers on a gold(111) surface. Biophys. J. 1999, 77, 964−972.

G

DOI: 10.1021/acs.langmuir.5b00859 Langmuir XXXX, XXX, XXX−XXX