In Situ STM and Vibrational Study of Nanometer-Scale Reorganization

Aug 1, 2017 - associated with the chemistry of DHPC by the aid of vibrational spectroscopy and the ... characterized as having hydrophilic and hydroph...
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In Situ STM and Vibrational Study of Nanometer-Scale Reorganization of a Phospholipid Monolayer Accompanied by Potential-Driven Headgroup Digestion Soichiro Matsunaga,† Hiroaki Shimizu,† 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 RIKEN, 2-1 Hirosawa, Wako, Saitama 351-0198, Japan § UMR 7213 CNRS, Faculté de Pharmacie, Université de Strasbourg, 74 route du Rhin, 67401 Illkirch, France ‡

ABSTRACT: In situ dynamic observation of model biological cell membranes, formed on a water/gold substrate interface, has been performed by the combination of electrochemical scanning tunneling microscopy and reflection infrared absorption vibrational spectroscopy. Monolayers of 1,2-dihexanoyl-sn-glycero-3-phosphocholine (DHPC) were formed on alkanethiol-modified gold surfaces in a buffer solution, and the microscopic phase transitions driven by electrochemical potential control were observed more in detail than our previous study on the same system [Electrochem. Commun. 2007, 9, 645−650]. This time the transitions were associated with the chemistry of DHPC by the aid of vibrational spectroscopy and the utilization of deuterium-labeled DHPC molecules. A negative potential shift solidifies the fluidic lipid layers into static striped or grainy features without notable chemical reactions. The first positive potential shift over the virginal DHPC monolayer breaks DHPC into choline and the corresponding phosphatidic acid (DHPA). This is the first case of a phospholipid electrochemical reaction microscopically detected at the solid surface.

1. INTRODUCTION Cell membranes are two-dimensional structures of phospholipid molecules. The physics and chemistry of these structures have been extensively investigated to understand biochemical reactions, such as metabolism, molecular recognition, and transportation across cell membranes.1 Moreover, the phospholipid layer is also recently considered as a potential material for nanometer-scale engineering.2 Phospholipid molecules involve several functional moieties within themselves, and the chemical reactions related to the membrane are sometimes modified and enhanced by forming heterogeneous aggregation of phospholipids,3 that is, phospholipid domains composed of different phospholipids or assembled into different molecular phases.4 To apply phospholipids for nanometer-scale engineering, the chemical reactions of phospholipid molecules should be controlled within the phospholipid layers, maintaining the basic membrane structures. Phospholipid molecules are characterized as having hydrophilic and hydrophobic parts in a single molecule. Within cell membranes, the hydrophilic parts are sticking out both into the cytoplasmic matrix and extracellular fluid, both of which are aqueous media. According to this structure of phospholipid membrane, one of the basic strategies for the engineering application is to desirably select, control, or exchange the hydrophilic moieties of phospholipid molecules assembled in the membrane.5 The © XXXX American Chemical Society

reaction to exchange the hydrophilic headgroup is called “transphosphatidylation”. Transphosphatidylation is in general achieved by enzymes, such as phospholipases.6,7 Although the enzyme reactions are highly selective with available concrete protocols, they are not always suitable for mass-scale production of modified phospholipids. As one possible solution for this, one can imagine electrochemical methods to split phospholipid molecules for the initial process of transphosphatidylation. Artificial electrolysis of phospholipid has never been considered as a route of transphosphatidylation. This imagination can be rationalized by considering the transmembrane potential of the real cell membrane. The transmembrane potential, namely the physiologically maintained electrochemical potential between the cytoplasmic matrix and the extracellular fluid, is one of the driving forces of the biochemical reactions and structural transformations on cell membranes.8 The transmembrane potential is mainly derived from ion balances between the inside and outside of cells, and the absolute value of the potential is about 70−200 mV.1 The potential imposes a strong electric field Received: June 7, 2017 Revised: July 30, 2017 Published: August 1, 2017 A

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C2H5OH solution for 1 day.21,22 This hydrophobic surface was fixed in the in situ STM cell. The desired phospholipid-containing solutions were poured to form the phospholipid monolayer on the hydrophobic substrate. As previously reported, 1.5 mM aqueous solution of DHPC was suitable for the monolayer formation.12 The DHPC molecules are anticipated to be soluble in water without aggregation, because the critical micelle concentration of DHPC is 15 mM.23,24 We obtained and stocked the phospholipids as CHCl3 solutions at −30 °C. To dispense DHPC, a portion of CHCl3 solution was taken into a small glass vial of the desired volume, and CHCl3 was evaporate by using a N2 gas. After drying completely, the calculated amount of buffer was poured into this vial, and it was vortexed and sonicated with an ultrasonic bath (US102, SND, Japan) for 10 min. 2.2.2. Preparation of Phospholipid Monolayer for in Situ IRAS. A multiple internal reflection (MIR) configuration was employed to enhance IR absorption signals from the phospholipid monolayer at the solution/substrate interface. For the attenuated total reflection (ATR) method,25 we used a silicon prism evaporated with a gold thin layer, on which 1-octanethiol SAM and a DHPC monolayer were deposited. Within the Si prism, the infrared beam with wavenumbers larger than 1200 cm−1 propagates without significant attenuation to achieve internal multiple reflection. Au thin films are necessary to embody a substrate to mimic the Au(111) single crystal, and at the same time it must be thin enough (99%), and partially deuterated DHPC of 1,2-dihexanoyl(d22)-sn-glycero-3-phosphocholine (Tail-DDHPC, purity >98%), 1,2-dihexanoyl-sn-glycero-3-phosphocholine1,1,2,2,-d4-N,N,N-trimethyl-d9 (Head-D-DHPC, purity >80%), and 1,2-dihexanoyl(d22)-sn-glycero-3-phosphocholine-1,1,2,2,-d4-N,N,Ntrimethyl-d9 (All-D-DHPC, purity >98%) were products in the form of CHCl3 solutions from Avanti Polar Lipids Inc. C2H5OH (spectroscopic grade), HClO4 (ultrapure grade), CHCl3 (>99%), NH3 (ultrapure grade), and 1-C8H17SH (>97%) were purchased from Kanto Chemicals, Japan. (3-Aminopropyl) trimethoxysilane (APTMS), purity >97%, was purchased from Sigma-Aldrich. Si prisms for in situ IRAS experiments were purchased from Pier optics, Japan. The Si prism ((111)-oriented, 20−100 Ω cm) size was cut by 45 mm × 15 mm × 2 mm. Both sides of the prism were mirror polished, and two edge faces were cut to 45° and were also mirror polished. The gold crystal was prepared from an Au wire, in diameter 1 mm and purity >99.999%, purchased from Furuya Metals, Japan. The electrodes and wirings for in situ STM and IRAS cell were made of Pt wires and sheets purchased from Nilaco, Japan. 2.2. Samples. 2.2.1. Preparation of Phospholipid Monolayer for in Situ STM. As a highly flat substrate for in situ STM, we used small (111) facets on a single-crystal gold bead19,20 covered with a 1-C8H17SH selfassembled monolayer (SAM) deposited by immersing into an 1 mM B

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Figure 1. STM image of Au deposited film on an APTMS-modified Si substrate in air. Bias voltage = 0.9 V (tip positive) and preset tunneling current = 1.0 nA. The graph shows the line profile along a line in the image. signal isolated by the aid of numerical normalization. Furthermore, we used DHPC reagents deuterated on the portions of our interest and observed the C−D signals isolated from the C−H signals from other portions. Figure 2 shows molecular structure of the partially deuterated DHPS reagents we used, which are called (A) normal DHPC, (B) headD-DHPC, (C) tail-D-DHPC, and (D) all-D-DHPC in this article. 2.3. Methods. 2.3.1. In Situ Scanning Tunneling Microscopy (in Situ STM). Figure 3A shows an overview of our experimental setup and sample of in situ STM measurement. Our STM setup was the model of Nanoscope E (Veeco instruments Inc.). The STM tips were commercial Pt/Ir (80:20) tips (Veeco). The tip surface was covered with a nail polish to minimize the Faraday current in the electrolyte. The measurement was performed in a homemade Teflon-PFA (PerFluoroAlkoxyethylene) cell, volume of liquid content = 2 mL, assembled with a Pt plate (6 mm × 20 mm) as the counter electrode. A pH-controlled NH4ClO4 buffer (0.05 M, pH 7.0 ± 0.1, prepared by mixing of NH3 and HClO4 aqueous solutions) was used as the supporting electrolyte. The potential of the sample was controlled by referring a quasi-reference Pt electrode which was measured to be 0.22 V vs Ag/AgCl in our present operational solution. Hereafter the electrode potential is indicated upon this reference potential. The potential of the STM tip was always adjusted at −0.4 V with respect to the sample substrate. The images were obtained in the constant current mode and the preset tunneling current was adjusted at a value between −0.6 and ∼−1.0 nA. The nature of the contrast of STM images is strongly related to the mechanism of tunneling current imaging. The tunneling is affected by not only the gap of tip and sample but also the electronic structure of the organic molecules. The mechanisms have been under discussion, and a part of them is mentioned in our previous report.14 2.3.2. In Situ Infrared Absorption Spectroscopy (in Situ IRAS). Figure 3B schematizes our experimental setup and sample for in situ IRAS. A Fourier transform infrared (FT-IR) spectrometer (Tensor, Bruker Optics) was used. A liquid-nitrogen-cooled HgCdTe detector was placed in the spectrometer setup. A potentiostat (#1110, Husou ElectroCmemical System, Japan) was connected to the electrodes directly to control the sample potential. The multiple internal reflections were achieved by setting the Si prism on four mirrors optics with an incident angle of 45°. From the geometry of the prism, approximately 20 times of reflections are expected. The ATR method not only enhances the weak IR absorption signal but also minimizes the background IR absorption of water. This optics was placed in the beam path within the spectrometer housing. The inside of the spectrometer was purged with a

Figure 2. Structural formula of phospholipids. (A) Normal DHPC (without deuteration), (B) head-D-DHPC, (C) tail-D-DHPC, (D) allD-DHPC, and (E) DHPA. continuous flow of dried nitrogen to remove background gases of CO2 and H2O in air. The Si prism was set in the measurement cell (volume of liquid content = 1.5 mL) made from Teflon-PFA, and assembled with a Pt plate (6 mm × 20 mm) as the counter electrode and a Pt wire as the reference electrode. The Si prism was pressed to Teflon O-ring over the liquid-tight compartment of the measurement cell. Two holes (∼1.5 mm in diameter) were made on the liquid-tight compartment for liquid exchange, which is necessary for switching isotope-labeled solutes. The detection volume of infrared beam upon each reflection at the Au surface is an important factor in discriminating the adsorbed species in the spectrum. The evanescent light penetrates into not only the phospholipid monolayer but also the buffer solution. The depth of penetration dp, defined as the exponential attenuation distance of the light electrical field amplitude, is given by

dp =

λ 2π n12 sin 2 θ − n2 2

(1)

where λ is the light wavelength in vacuum, n1 and n2 are the refractive indices of Si and water, respectively, and θ is the angle of light incidence from Si bulk to Si surface.25 For the monochromatic light of 2300 cm−1, which is the median of our infrared spectral range, the values of n1 and n2 are 3.4429 and 1.3530 at room temperature, respectively. With θ = 45°, dp is calculated to be 0.34 μm, neglecting the Au layer. Suppose one DHPC single molecule occupies 0.52 nm2 in a monolayer,31 1.9 × 106 DHPCs are involved in a 1 μm × 1 μm square area on the substrate. On the other hand in the solution above this area, C

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Figure 4. Comparison of in situ IR spectra (C−D stretching region, 2300−2000 cm−1) of all-D-DHPC monolayer with and without all-DDHPC molecules in the bulk solution. The black curve was recorded on a 1-octanethiopl/Au(111) substrate after incubating for 3 h in all-DDHPC-containing (1.5 mM) NH4ClO4 solution (0.05 mM, pH 7.0). Then, the all-D-DHPC solution was replaced with pure NH4ClO4 solution (0.05 mM, pH = 7.0). The red line was recorded after 10 h of this solution replacement.

Figure 3. Schematics of our experimental apparatuses and sample settings of (A) in situ STM and (B) in situ IRAS. the light penetrates within the span of dp = 0.34 μm; hence, the number of solute DHPCs is 1 μm × 1 μm × 0.34 μm × 1.5 mmol L−1 = 3.1 × 105. On the basis of this estimation, approximately 85% of the infrared detected DHPCs are adsorbed on the interface, and the rest of the 15% are dissolved in the bulk solutions. This estimation was actually verified by measuring the intensities of IRAS peaks by exchanging the buffer solution. First, the liquid cell was filled with 1.5 mM DHPC dissolved in 0.05 M aqueous NH4ClO4 solution, and a spectrum was recorded. This spectrum involves the DHPC signal from both the monolayer on the substrate and the molecules in the buffer solution. Then we exchanged the solution with 5 times volume of pure 0.05 M NH4ClO4 solution to remove the DHPC dissolved in the solution. As shown in Figure 4, after exchanging the solution, the signal intensity from DHPC molecules was decreased to approximately 70% of the initial intensity, and this is reasonable as estimated above. If a DHPC molecule in the lipid layer absorbs IR more than a DHPC molecule in the bulk solution for some reasons, such as by the mechanism of surface-enhanced infrared absorption (SEIRA),32,33 the signal intensity from the lipid layer will be stronger. We additionally evaluated the effect of SEIRA on the present Au/APTMS/Si(111) substrate. For this, the 1-octyl monolayer directly bonded to the Si(111) prism surface was prepared by 1-octylmagnesium chloride Grignard reaction.34 This substrate and the present 1-octanethiol-modified Au/ APTMS/Si(111) were equally immersed in the DHPC solution, and in situ IRAS was regularly recorded. Figure 5 shows the spectra obtained from DHPC spread on these two substrates. In the range, strong absorption of the CO stretching mode appears around 1730 cm−1. The intensity with the Au layer is approximately ten times larger than without Au. Preferably, SEIRA seems contributing selective detection for DHPC adsorbates. The originally reported SEIRA exhibits more than hundreds of times of enhancement.32 In those works the Au films were composed of particles with diameters of a few tens of nanometers and similar heights,32 whereas our Au substrates were planar as shown in Figure 1 and seem not so optimal for SEIRA.

Figure 5. Comparison of in situ IR spectra (CO stretching region, 1850−1650 cm−1) of DHPC spread on 1-octanethiol/Au/APTEM/Si (black) and octyl-modified Si (red).

The intensity of the spectrum was not altered at least 4 days after the lipid−containing solution was replaced with pure NH4ClO4. This indicates that the DHPC monolayer was stably maintained for an extended period of time on the substrate without desorbing into the bulk solution. This stability also means that the exchanging speed between the lipid in solution and that on the substrate is very slow, and therefore we can interpret in situ STM and IR results without considering the exchange of the lipid between the solution and the substrate.

3. RESULTS 3.1. In Situ STM Observation of the Phospholipid Monolayer. Previously, we showed nanometer scale structural changes of DHPC monolayer induced by the electrochemically potential shift of the substrate. In the present work, we extended this experiment and studied the molecular mechanism that drives the structure alteration. The sequence of the potential control and representative monolayer structures are shown in Figure 6. The DHPC monolayer was formed on 1-octanethiol SAM substrate at 0.0 V, as a fluidic DHPC monolayer (“Fluidic I”, Figure 6A). The fluidic character was apparent in the motion of sliding “lipid windows” as a vacancy of DHPC monolayer, and the STM observed height of the DHPC monolayer was 0.19 ± 0.02 nm, which reflects not only the physical height but also the electron conductivity of the layer. Then, the substrate potential was swept down to −0.2 V, and after 17.5 h, a striped structure was observed (“striped”, Figure 6B). The average interval of the stripes was 4.3 ± 1.3 nm, and the D

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Figure 6. Series of in situ STM images of the DHPC monolayer on the 1-octanethiopl/Au(111) substrate. The concentration of DHPC was 1.5 mM in 0.05 mM NH4ClO4 solution (pH 7.0). According to the potential shifting indicated in the lower viewgraph, (A) fluidic I, (B) striped, (C) fluidic II, and (D) grainy appeared sequentially. Panels B−D were recorded at 17, 6.5, and 13.5 h after the potential shifting. At time (C′) and (D′), STM images similar to (C) and (D), respectively, were observed. The tunneling current was kept at −0.6 or −1.0 nA.

Figure 7. Sequence of in situ STM images in the conversion from grainy to fluidic II. The concentration of DHPC was 1.5 mM in 0.05 mM NH4ClO4 solution (pH 7.0). The hour number on the top of each frame indicates the time after the negative potential shift from +0.20 to −0.20 V. The tunneling current was kept at −1.0 nA.

Figure 8. In situ STM images of the DHPC monolayer on the 1-octanethiopl/Au(111) substrate. The concentration of the lipid was 2 mM in 0.05 mM NH4ClO4 solution (pH 7.0). The bottom graph shows the potential shifting profile. Indexes in the STM images correspond to those in the graph. Panels B, A, and B′ were recorded at 10, 8, and 11 h after the potential shifting. Fluidic I and striped were reversibly observed. The tunneling current was kept at −1.0 nA.

height was 0.33 ± 0.11 nm. The stripes covered a whole area of surface and override the monatomic steps of the gold substrate. By stepping up and keeping the potential at +0.2 V for 6.5 h, a fluidic feature was again observed (Figure 6C). However, the average thickness of the lipid layer was 0.22 ± 0.03 nm, slightly but definitely larger than fluidic I seen in the very first cycle, and we distinguish this fluidic layer by designating as “fluidic II”.

Sweeping back the potential to −0.2 V and within a period of 13.5 h, a slow propagation of solid structure was observed. Many grains were observed as shown in Figure 6D, and the grains covered the whole area of the substrate. The grains were elliptic, and the average size was 13.3 ± 1.4 nm × 4.6 ± 0.6 nm with a height of 0.7 ± 0.1 nm. We named this structure “grainy”. E

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Langmuir In 21.5 h after stepping the substrate potential to +0.2 V, fluidic II phase was restored. By further potential cycling between +0.2 and −0.2 V, just Fluidic II and grainy appeared sequentially. Figure 7 shows the slow unbundling process of grainy into fluidic II after stepping the potential to +0.2 V serially imaged after 21.5 h at the latest. The grains merge together toward formation of a flat layer. This time we tried to clarify the electrochemical potential ranges of DHPC fluidization (fluidic I and II) and solidification (stripe and grainy). We limited the potential range by stepping between 0 and −0.2 V and observed the phase transition. Figure 8 shows the potential program and resulting STM images. Fluidic I and striped were reversibly observed after the indicated periods at 0.0 and −0.2 V, respectively. Fluidic II and grainy were never observed in this sequence. It seems that once these phases appeared after increasing the potential to +0.2 V, the monolayer does not exhibit fluidic I and striped any more. Thus, changing to fluidic II and grainy is an irreversible process. There seems to be an irreversible process for DHPC upon setting the potential at +0.2 V for the first time. In Figure 9, we employed 1,2-didecanoyl-sn-phosphocholine (DDPC), instead of DHPC, spread over 1-dodecanethiol-SAM-

recorded more than 8 h after each of the potential steps. The IRAS spectra are nominated according to the structural changeover among fluidic I, striped, fluidic II, and grainy, by assuming that the DHPC layer morphology changeover on the Au(111) film is all of the same as that observed by STM on the Au(111) bead crystals as seen in Figure 6. The partially deuterated DHPC molecules, head-D-DHPC and tail-D-DHPC shown in Figure 2, were utilized for this purpose. Table 1 shows the vibrational modes and wavenumbers Table 1. Assignment of Deuterated Methyl (CD3) and Methylene (CD2) Stretching Mode in Choline Part and Fatty Acid Part in Phosphocholine34

a

wavenumber (cm−1)

modea

2275 2231 2120 2217 2195 2105 2071

CD3 as (choline) CD2 as (choline) CD2 s (choline) CD3 as (fatty acid) CD2 as (fatty acid) CD3 s (fatty acid) CD2 s (fatty acid)

as: asymmetric stretching mode, s: symmetric stretching mode.

of deuterated methyl and methylene parts, according to Gauger’s experimental results35 on a partially deuterated 1,2-dimyristoylsn-glycero-3-phosphocholine multilayer film in dried air. Within the vibrational spectra, we focused on the C−D stretching region (from 2000 to 2300 cm−1), CO stretching region (around 1730 cm−1), and the phosphate stretching region (from 1200 to 1300 cm−1). For a better understanding of the relationship between the spectrum and the structure at each potential, hereafter we label the spectra according to the structure observed in STM images seen in the same cycles of potential, that is, fluidic I, striped, fluidic II, and grainy. 3.2.1. Spectra of Tail-D-DHPC. Figure 10 shows the IR absorption spectra of the tail-D-DHPC monolayer between 1650 and 2300 cm−1, involving the C−D stretching signals from tailD-DHPC monolayer. The largest peak at 1730 cm−1 is assigned to the CO stretching mode in the ester part of the hexanoyl group. The three peaks and one shoulder peak around 2300 cm−1 are associated with C−D stretching vibration in CD2 groups or

Figure 9. In situ STM images of the DHPC (6:0 PC) monolayer on the 1-octanthiol/Au(111) substrate and the DDPC (10:0 PC) monolayer on the 1-dodecanthiol/Au(111). Both images were obtained after more than 8 h of changing the substrate potential from 0 to −0.2 V. The concentration of the lipid was 1.5 mM for each, in 0.05 mM NH4ClO4 solution (pH 7.0). The tunneling current was kept at −1.0 nA.

modified Au(111). Figure 9 shows in situ STM images of the DHPC/1-octanethiol and DDPC/1-dodecanthiol systems after the substrate the potential was first swept to −0.2 V. “Striped” structure was observed also in the DDPC/1-dodecanethiol/ Au(111) system after 8 h. 3.2. In Situ IRAS of the Phospholipid Monolayer. In situ IRAS measurement was purposed to pursue the distribution of whole DHPC molecules in the adlayer and solution, as well as chemical reactions of the DHPC molecules. A certain alteration of DHPC can be associated with the above-mentioned irreversible transition from striped to fluidic II in the very first positive potential sweep. The potential cycling sequence was the same as that in STM observation. The sequence was composed of starting from 0 V, stepping down to −0.2 V for more than 21 h, stepping up to +0.20 V for more than 20 h, and repetition of this cycle. IRAS was

Figure 10. Sequence of in situ IR absorption spectra of the tail-D-DHPC monolayer on the 1-octanthiol/Au(111) substrate. The concentration of the lipid was 2 mM in 0.05 mM NH4ClO4 solution (pH 7.0). The spectra were recorded more than 8 h after each of the potential steps. The spectral nominations “fluidic I”, “striped”, “fluidic II”, and “grainy” are made in accordance with the STM features observed in potential cycling process among 0, −0.2, and +0.2 V. F

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In order to understand the change of peaks in each step, each spectrum was subtracted by the spectrum recorded in the step immediately before. The spectra subtracted in this way are shown in Figure 13, converted from Figure 12. In Figure 13, only the

CD3 groups in the fatty acid parts, respectively, as shown in Table 1. Positions of these five signals did not change in the whole course of the structural changes and the intensities of the peaks were maintained within 0.1% deviation from the initial intensity. The invariance tail-D-DHPC spectra tell that the fatty acid chains of DHPC were not involved in the structural changes observed by STM. 3.2.2. Spectra of Head-D-DHPC. Figure 11 shows the spectra for a head-D-DHPC monolayer between 1650 and 2300 cm−1,

Figure 13. Numerically calculated spectra by step-by-step subtraction on the basis of Figure 12. In the top column, the colors of curves are defined by the spectra of divided operand/the spectra of dividing operand.

black solid line is just a duplication of fluidic I of head-D-DHPC. Spectrum “striped-fluidic I” is the spectrum of striped subtracted by the spectrum of fluidic I shown in Figure 12. Spectrum “striped-fluidic I” is flat, indicating that the choline group remained unreacted between fluidic I and striped. In the next spectrum “fluidic II-striped”, the polarity of the CD2 symmetric stretching peak is inversed, demonstrating a decrease in the CD2 symmetric stretching intensity. In the following spectrum “grainy-fluidic II”, the peak returns to the regular polarity. This series of spectra indicates that the choline part represents the chemistry of DHPC under the potential stepping. In the following repetitive cycling, every “fluidic II(2)-grainy” and “grainy(2)-fluidic II(2)” spectrum the CD2 peak is in the inverted polarity and the peak heights of “fluidic II(2)-grainy” and “grainy(2)-fluidic II(2)” are gradually weakened cycle by cycle. 3.2.3. Spectra of the Phosphate Part. Figure 14 shows the spectra of the all-D-DHPC monolayer between 1220 and 1280 cm−1. This region is for the vibrations of the phosphate parts. In all of fluidic I, striped, fluidic II, and grainy, the phosphate part clearly changed. In fluidic I and striped, three peaks were observed at 1233, 1259, and 1278 cm−1. In fluidic II and grainy, two peaks are observed at 1230 and 1280 cm−1. The intensity

Figure 11. Sequence of in situ IR absorption spectra of the head-DDHPC monolayer on the 1-octanthiol/Au(111) substrate. The concentration of the lipid was 2 mM in 0.05 mM NH4ClO4 solution (pH 7.0). The spectra were recorded more than 8 h after each of the potential steps.

involving the C−D stretching signals in the course of potential cycling. The largest peak at 1730 cm−1 is assigned to the CO stretching mode in the ester part of fatty acids; all spectra are the same as in Figure 10. The peaks around 2100 cm−1 are associated with C−D stretching vibration in CD2 groups and CD3 groups in the choline group. The CO stretching peak at 1730 cm−1 was invariant in the whole course of potential cycling. This is again the same as the tail-D-DHPC experiment. The point-of-interest region from 2000 to 2300 cm−1 in Figure 11 is enlarged and shown in Figure 12. The peak at 2120 cm−1 is

Figure 12. Spectral region of CD2 stretching, drawn by expanding the graph of Figure 11 from 2050 to 2250 cm−1.

assigned to the symmetric stretching motion of the CD2 groups in the choline group. The stretching modes of the CD3 group were practically undetectable in our experiments. As the potential was cycled, the peak of the CD2 asymmetric stretching mode shifted, and the intensity of the peak is slightly but definitely changed.

Figure 14. Sequence of in situ IR absorption spectra of All-D-DHPC monolayer on the substrate. Concentration of the lipid was 2 mM in 0.05 mM NH4ClO4 solution (pH 7.0). Peaks derived from P−O bonds in phospholipid appear in this region. The spectrum were recorded more than 8 h after each of the potential steps. G

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Figure 15. Schematics of DHPC morphologies that appeared as fluidic I, striped, fluidic II, and grainy upon the electrode potential shifting.

cm−1 was invariantly observed in all spectra of A−D in Figure 14. It is seen that the P−O bond at 1278 cm−1 is shared in all four phases. By referring to the intensity of the CO stretching peak described in the previous part, it is clearly shown that DHPC was maintained on the substrate, and their amphiphilic property was also maintained throughout the four structural changes. The glycerol-bound P−O was necessary for the amphiphilic structure of the phospholipid as shown in Figure 2. Therefore, the 1278 cm−1 peak should be related with glycerol-bound P−O. The peak at 1260 cm−1 is a characteristic P−O peak derived phosphocholine.41,42 The other 1233 cm−1 peak is consequently assigned to the nonbind P−O (conjugation of PO and P−O−).

ratio of these peaks is not the same. This indicates that the phosphate part is associated with the structural change of the DHPC monolayer. In Figure 14, the spectra for the second cycles between fluidic II and grainy are also shown. They are tagged with (2) in Figure 14. The first spectra of fluidic II and grainy are very similar to the second ones. The frequencies of P−O stretching vibration modes in the phosphate group are sensitive to the headgroup substituent, such as choline, ethanolamine, L-serine, and so on,36−39 and moreover the P−O peaks shift depending on the degree of hydration. Therefore, it is difficult to assign vibrational peak position to the incorporated bonds. Anyway the change of the peaks depending on the structure in our experiments and understanding of origins of the peaks will give us more information on the molecular structure in each structure. On the basis of the IR result in the P−O stretching region, we tried to assign the peak origins. In (A) fluidic I spectrum, three peaks of phosphate are positioned at 1233, 1259, and 1278 cm−1. Intact DHPC molecules have the three kinds of P−O bonds in the phosphate part, namely, the glycerol-bound, the cholinebound, and two equivalent terminal bonds, which are the conjugation of PO and P−O− (see Figure 2A). These three separated peaks can be assigned to these three kinds of P−O bonds. In a previous report,40 the three peaks were assigned as the O− P−O asymmetric vibrations which shift depending on the degree of hydration. As the phosphate part is more hydrated, the vibrational peak shifts lower. In the present case, the peak at 1278

4. DISCUSSION In our previous report,12 we showed in situ STM images in a single potential sequence and speculatively discussed the mechanism behind the alternations. In the present report, we confirmed a certain generality of the transitions observed for DHPC and DDPC by STM. The transition potential from striped to fluidic (II) was also clarified by STM. On the basis of those facts revealed by in situ STM, we performed an IRAS study in parallel to understand precisely the microscopic reaction mechanism behind the phospholipid monolayer under the potential control. Figure 15 shows schematic drawings of DHPC molecules on the substrate in fluidic I, striped, fluidic II, and grainy. H

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that the driving force of the grain aggregation is electrostatic interactions between negatively charged DHPA and cations in solution, such as H+, NH4+, H3O+, and choline+, attracted by the negative electrode potential.44 A seen in Figure 14, the number and the shapes of the IR peaks of the phosphate part for fluidic II would not change the first, second, and following cycles, passing over the alteration reaction between fluidic II and grainy. The integration of fluidic II phase is guaranteed by the hydrophobic force of the DHPA fatty acid part conforming to the substrate covered with 1-octanethiol. According to Figure 14, the phosphate peaks exhibited completely reversible changes, whereas in Figures 12 and 13 the CD2 peak in the choline part regains in every step of fluidic II to grainy, but the CD2 peak in the next grainy phase is weakened. Some of the detached choline molecules from DHPC stay near the DHPA monolayer but some of them diffuse into the bulk solution when fluidic II is maintained. Finally, we discuss a mechanism behind choline digestion from DHPC at the anodic potential. Detachment of choline from PC is understood as hydrolysis of phosphoester. We believe that the hydrolysis is enhanced by hydroxide ions attracted by the anodic potential toward the substrate. One single DHPC molecule has two types of ester bonds, that is, the two glycerol ester bonds to hexyl tail parts and one phosphoester bond to the choline head part. Our IR experiment indicates that the phosphoester part was selectively hydrolyzed. This selective hydrolysis might be due to the steric effect for the OH− attacking DHPC from the bulk solution. As seen in the STM images of fluidic I, the mobile DHPC monolayer was flat. The DHPC molecules in the monolayer must be closely packed with their head groups in contact with the solution. Then the choline phosphoester bonds are closer to the bulk solution, and the pairs of glycerol ester bonds are located in a deeper position. The probability of hydrolyzing reagents reaching phosphoester bonds might be naturally higher than that to the glycerol ester bonds. A similar steric effect on the phospholipid layer was reported in a system of selective binding of small peptides to phosphatidylethanolamine.45

At 0 V in the starting stage, in situ STM indicates that DHPC molecules form a fluidic monolayer (“fluidic I”) and IR spectroscopic data indicates that fluidic I contains intact DHPC molecules. According to the invariance of the CO stretching peak in Figures 10 and 11, the DHPC monolayer is stably maintained on the substrate in the experiments, because the position and intensity of their CO vibrational peak were maintained for several tens of hours in the IR experiments (Figures 10 and 11). STM observation discerns the striped structure formed in cathodic potential application as a solid adlayer of DHPC molecules two-dimensionally arrayed within uniformly oriented islands. As long as the potential is set below 0 V, striped structure is resumed by sweeping back to −0.2 V. The IR spectrum of the striped showed no difference from that of fluidic I both in choline and fatty acid parts. The signal in the phosphate part certainly perceivably varies, but three kinds of P−O vibrational signals survive. These IR results indicate that DHPC molecules stay unchanged in striped and fluidic I. The small difference of the phosphate part indicates the phosphate part was related with the orientation of DHPC. A similar ordered-structure of phosphocholine was reported in the case of the 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) layer directly spread on Au(111) substrate by Lipkowski et al.,43 although the DMPC molecules seem arrayed under the influence of a bare metallic substrate. They considered a hemimicellar aggregation of the phospholipid molecules for the ordered structure. In the case of striped DHPC, the distance between the two nearest stripes, stripe height, and length of the wavy pattern on a stripe are close to those of the DMPC hemimicellar structures.43 It is notable that the alignment of fluidic I molecules into striped structure is reversibly stimulated by the potential reversing. It was already noted that the monolayer heights of fluidic I and II were 0.19 and 0.22 nm, respectively, and distinctively different from each other. As shown in the IR spectral region in the phosphate part (Figure 14, 1200−1300 cm−1), we can recognize three peaks for fluidic I and just two peaks for fluidic II. The peak at 1260 cm−1, assigned to the P−O group binding to the choline group in the DHPC molecule, is missing in fluidic II, and 1233 cm−1 peak became stronger. The loss of the peak at 1260 cm−1 indicates detachment of the choline group from DHPC, resulting in the formation of 1,2-dihexanoyl-sn-glycero-3-phosphate (DHPA). DHPA is actually composed of the fluidic II phase in the STM images. The molecular structures of DHPC and DHPA are shown in Figure 15. A DHPA molecule includes one glycerol-bound P−O bond and three terminal P−O bonds which are in conjugation with PO and P−O−. DHPC and DHPA are differentiated by the presence and absence of the choline-bound P−O bond. As shown in Figures 12 and 13, the CD2 peak intensity of choline in head-D-DHPC becomes weaker in fluidic II, indicating that the choline part is detached from DHPC in the fluidic II. The choline CD2 peak did not completely disappear in the spectrum as shown in Figure 12. Some of the detached choline molecules remained near the substrate within the detection volume of the present IR-ATR sample system. The existence of the detached choline part will be discussed again later. When the substrate potential was swept to cathodic again, grainy images were observed. The grains are sized 13.3 nm × 4.6 nm with a height of 0.7 ± 0.1 nm. The grains are naturally considered to be aggregates of DHPA molecules. We consider

5. CONCLUSION We performed in situ STM and IRAS observation on a fluidic 1,2dihexanoyl-sn-glycero-3-phosphocholine (DHPC)/1-octanethiol SAM on Au(111) substrate in aqueous solution. By IRAS vibrational analysis, we pursued the chemistry of DHPC accompanied by the structural alteration upon electrochemical potential shifting among fluidic I, striped, fluidic II, and grainy observed by in situ STM. By the aid of partly deuteriumlabeled DHPC molecules, the IRAS analysis demonstrated that fluidic I and striped are composed of intact DHPC and that scission of the choline phosphoester bond takes place upon the very first application of anodic potential, resulting in the lipid layer being composed of DHPA. The integrity of fluidic phospholipid layers, the position of bond scission in the DHPC molecule, and the mechanism behind phase changes observed by STM are discussed in detail. This work verifies a notable applicability of this combination of nanoscopic and vibrational analysis for biological entities immersed in aqueous solution. To our knowledge, this is the first case of analyzing the bond breaking within a rather largesized molecule, such as a phospholipid, within a molecular monolayer composed over a solid surface. Not only in the discipline of biology but also in general surface science, I

DOI: 10.1021/acs.langmuir.7b01912 Langmuir XXXX, XXX, XXX−XXX

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application of highly sensitive multitechnique analysis for complex systems of materials is looked forward to.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Soichiro Matsunaga: 0000-0002-5003-9566 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This study was financially supported in part by RIKEN President Discretionary Fund (2004−2006, 2010−2012), Lipid Dynamics Program, and Integrated Lipidology Program of RIKEN and Grants-in-Aid for Scientific Research on 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 the support from Research Fellowships for Young Scientists of the Japan Society for the Promotion of Science.

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K

DOI: 10.1021/acs.langmuir.7b01912 Langmuir XXXX, XXX, XXX−XXX