Morphology and Physical Properties of Hydrophilic-Polymer-Modified

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Biological and Environmental Phenomena at the Interface

Morphology and Physical Properties of HydrophilicPolymer-Modified Lipids in Supported Lipid Bilayers Yasuhiro Kakimoto, Yoshihiro Tachihara, Yoshiaki Okamoto, Keisuke Miyazawa, Takeshi Fukuma, and Ryugo Tero Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b00870 • Publication Date (Web): 23 May 2018 Downloaded from http://pubs.acs.org on May 23, 2018

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Morphology and Physical Properties of HydrophilicPolymer-Modified Lipids in Supported Lipid Bilayers Yasuhiro Kakimoto1, Yoshihiro Tachihara1, Yoshiaki Okamoto1, Keisuke Miyazawa2, Takeshi Fukuma2, 3 and Ryugo Tero1,* 1. Department of Environmental and Life Sciences, Toyohashi University of Technology, Toyohashi, Aichi 441-8580, Japan. 2. Division of Electrical Engineering and Computer Science, Kanazawa University, Kakumamachi, Kanazawa 920-1192, Japan. 3. Nano Life Science Institute (WPI-NanoLSI), Kakuma-machi, Kanazawa 920-1192, Japan.

1 ACS Paragon Plus Environment

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ABSTRACT: Lipid molecules such as glycolipids which are modified with hydrophilic biopolymers participate in the biochemical reactions occurring on cell membranes. Their functions and efficiency are determined by the formation of microdomains and their physical properties. We investigated the morphology and properties of domains induced by the hydrophilic-polymermodified lipid applying poly-ethylene glycol (PEG)-modified lipid as a model modified lipid. We formed supported lipid bilayers (SLBs) using a 0–10 mol % range of PEG-modified lipid concentration (CPEG). We studied its morphology and fluidity by fluorescence microscopy, the fluorescence recovery after photobleaching (FRAP) method and atomic force microscopy (AFM). Fluorescence images showed that domains rich in PEG-modified lipid appeared and SLB fluidity decreased when CPEG ≥ 5 %. AFM topographies showed that clusters of the PEG-modified lipid appeared prior to domain formation and the PEG-lipid-rich domains were observed as depressions. Frequency-modulation AFM revealed a force-dependent appearance of the PEG-lipid-rich domain.


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Introduction Cell membranes are the outermost layers of cells. They are reaction fields recognizing the ambient environment and transporting information and materials.1,2 The basic structure of the cell membrane is a bimolecular sheet of amphiphilic lipid molecules. All natural cell membranes contain lipids modified with hydrophilic carbohydrates (glycans) such as glycolipids. Lipopolysaccharide, a gram-negative bacterial glycolipid, is found in the outer leaflet of the outer membrane. It confers cellular immunity via toll-like receptor 4.3,4 Gangliosides participate in various neuronal and brain cell functions such as signal transduction and cellular recognition.5–7 Clustering and domain formation in these molecules are key factors in cell membrane recognition and transport. This mechanism is represented by the “raft” concept.8,9 Lipid aggregation also affects physical properties and chemical functionalities and modulates the cell membrane reaction.10 Interactions within the hydrophobic core of the lipid bilayer strongly affect localization and domain formation of lipids and membrane proteins. Typical examples are the phase separations of the gel and liquid crystalline phases, and between the liquid ordered and liquid disordered phases in artificial lipid bilayer systems.11 The driving forces of these lipid localizations are the thermal phase transition of hydrophobic acyl chains, and cholesterol partition in saturated acyl chains, respectively. However, the large head groups of lipids with hydrophilic moieties also induce domain .6,12,13 This type of domain formation is poorly understood compared to those induced by hydrophobic interactions. In the former, competition occurs between attractive and repulsive interactions because of hydrogen bonding, hydration, and thermal fluctuation. Glycolipid aggregation and domain formation in lipid bilayer membranes have been investigated,12,14–18 and experimental platforms and methodologies are required. Lipid molecules


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with head groups modified by synthetic hydrophilic polymers such as polyethylene glycol (PEG) have been applied in design and control of biointerfaces.19–25 The aggregation states and molecular orientations of PEG-lipids such as the mushroom and brush regimes strongly affect their function and biocompatibility.24–31 Therefore behavior of hydrophilic-polymer-modified lipids is an important research subject in both native and artificial biomembrane systems. Supported lipid bilayers (SLBs) are artificial lipid bilayer membranes situated at solid-liquid interfaces.32–36 They are used to investigate the fundamental physicochemical properties and molecular distribution of lipid bilayers with atomic force microscope (AFM) and fluorescencemicroscope-based methods.36–46 In this study, we examined the morphology and membrane fluidity of the SLB containing PEG-modified lipid (PEG-SLB) using a fluorescence microscope and AFM. The objective was to determine the influence of a large hydrophilic head (rather than a hydrophobic tail) on lipid localization in a lipid bilayer and its effects on its physical properties. Hydrophilic polymer interactions induce clusters and domains in lipid bilayer membranes but they also complicate AFM topography because of the artificial forces they create between a sample and a cantilever tip. These experimental methods can be applied to the study of glycolipids and other natural-source lipids with hydrophilic polymers.

Materials and methods Preparation of lipid vesicle suspensions and supported lipid bilayers 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC), 1,2-distearoyl-sn-glycero-3phosphoethanolamine-N-[amino (polyethylene glycol) 2000] (PEG-DSPE), 1,2-dipalmitoyl-sn-


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glycero-3-phosphoethanolamine-N-lissamine rhodamine B (Rb-DPPE, Ex/Em : 560/583 nm), and 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[poly (ethylene glycol) 2000-N'carboxyfluorescein] (CF-PEG-DSPE, Ex/Em : 485/523 nm) were purchased from Avanti Polar Lipids, Inc. (Alabaster, AL, USA), and used without purification. Chloroform solutions of DOPC, PEG-DSPE (0-10 mol%), and a dye-labeled lipid (0.5 mol%) were mixed in a glass vial. Rb-DPPE was used as a fluorescence probe unless otherwise noted. The solution was dried with a nitrogen gas stream followed by overnight evacuation. A buffer solution (100 mM KCl and 25 mM HEPES/NaOH: pH 7.4) was added to the vacuum-dried lipid mixture film. A lipid vesicle suspension was then prepared by vortexing at 45 °C, freeze-thawing through five cycles, and extruding through a 100-nm polycarbonate filter. SLB was prepared by the vesicle fusion method on a thermally oxidized SiO2/Si substrate precleaned by boiling in piranha solution (3:1 v/v mixture of conc. H2SO4 and 30% H2O2 aqueous solution).44,47 The SiO2/Si substrate was incubated in the extruded vesicle suspension for 60-90 min at 45 °C. After the incubation, the excess vesicles in the liquid phase were washed out by serial exchanges of the vesicle suspension with buffer solution. Fluorescence microscope observation An epi-fluorescence microscope (BX51WI, Olympus, Tokyo, Japan) equipped with a 60× water-immersion lens (LUMPlan FL 60×, NA = 1.00) was used for wide-field fluorescence and qualitative fluorescence recovery after photobleaching (FRAP) observations at 25 °C. The spatial resolution at the wavelength of 580 nm was approximately 350 nm based on the Rayleigh criterion. A laser scanning microscope (A1, Nikon, Tokyo, Japan) was used for the quantitative diffusion coefficient (D) and mobile fraction (Fm) measurements by the FRAP method at


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25 °C.48–50 The experimental conditions were the same as those described in a previous study.51 Briefly, photobleaching was performed on a 26.2 µm × 26.2 µm square for 2 s using excitation light ~3500 times brighter than that used for observation. D and Fm were obtained by fitting the fractional recovery time course (f(t)) with the equation of Soumpasis and Berquand:49,50 exp

(1)

where I0 and I1 are modified Bessel functions of the first kind on the order of 0 and 1, respectively; τD = w2/(4D) is the characteristic diffusion time; w is the radius of the bleached region; and D is the diffusion coefficient. Five FRAP curves were measured at different positions of each sample to obtain average D and Fm. Atomic force microscope observation AFM observation was performed using the PicoPlus 5500 (Keysight Technologies, Inc., Santa Rosa, CA, USA, formerly Molecular Imaging, Corp.) in the amplitude-modulation mode (AMAFM) in the buffer solution at 25 °C. A cantilever with a spring constant of 0.09 N/m (resonant frequency (fres): 110 kHz; BL-AC40TS-C2, Olympus, Tokyo, Japan) was used. Topography, phase-shift image, and amplitude image were obtained simultaneously with 256 × 256 pixels. Frequency-modulation AFM (FM-AFM) observation was performed with a homemade ultralownoise FM-AFM instrument.52,53 The cantilever was either NCH-AuD (fres: 150 kHz in water; spring constant: 40 N/m; Nanoworld, Neuchâtel, Switzerland) or AC55 (fres: 1 MHz in water; spring constant: 85 N/m; Olympus, Tokyo, Japan).


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Results Macroscopic morphology observed by fluorescence microscopy Figure 1 shows fluorescence images of the PEG-SLBs with a 0–10 % PEG-DSPE concentration (CPEG) range. Uniform fluorescence intensity was obtained from the PEG-SLB with CPEG = 0 % and 2.5 % (Figures 1a and 1b). Figures 1c and 1d show the fluorescence images of the PEG-SLB with CPEG = 5 % containing both Rb-DPPE (red) and CF-PEG-DSPE (green). They were obtained at the same position using the mirror unit for Rb (Figure 1c) and CF (Figure 1d). In the image of Rb-DPPE (Figure 1c) PEG-SLB had dark regions that were poor in RbDPPE. The image of CF-PEG-DSPE (Figure 1d) shows brighter regions than the surrounding that had a uniform fluorescence intensity. The dark regions in Figure 1c and the bright regions in Figure 1d overlapped perfectly in the merged image (Figure 1e, represented by a white arrow). Therefore, the dark regions in Figure 1c are not defects but rather domains rich in PEG-DSPE. A defect, at which lipid bilayer is absent, is observed dark both in the Rb-DPPE and CF-PEGDPPE images, as indicated by the black arrow in Figure 1e. Figures 1c–1d show that PEGDSPE-rich domain emerged in the uniform SLB as CPEG increased to 5 %. Rb-DPPE was excluded from the PEG-DSPE-domain and existed mainly in the surrounding region, whereas CF-PEG-DPPE preferentially located in the PEG-DSPE domain but was still distributed in the surrounding region (uniform fluorescence intensities in Figure 1d). The number density of the domain was 8.0 × 10-3 μm-2. At CPEG = 10 % (Figure 1f), only a few dark domains can be clearly discerned but the fluorescence intensity was heterogeneous in most of the SLB. Generally only a single bilayer is formed by the vesicle fusion method using small unilamellar vesicles.36 Layer number is distinguished from the fluorescence intensity if multiple bilayer membranes form.54,55 All the SLBs in this study consisted of a single bilayer membrane.


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Figure 1. Fluorescence images of the PEG-SLBs with CPEG = 0–10 %. (a, b) PEG-SLB with (a) CPEG = 0 % (DOPC-SLB) and (b) CPEG = 2.5 % labeled with Rb-DPPE (Ex/Em: 560/583 nm), (c, d) PEG-SLB with CPEG = 5 % labeled with Rb-DPPE and CF-PEG-DSPE (Ex/Em: 485/523 nm), (c) Rb-DPPE and (d) CF-PEG-DSPE images obtained at the same position. (e) Merged image of (c) and (d). Insert: magnified image of the dotted square region. Representative PEG-DSPE-rich green regions are indicated with a white arrow. Dark defect is indicated by a black arrow. (f) PEG-SLB with CPEG = 10 % labeled with Rb-DPPE. Fluidity and mobile fraction Figure 2 shows the FRAP process of PEG-SLBs with CPEG = 0–10 %. The fluorescence intensity of PEG-SLBs with CPEG = 0–5 % recovered over time (Figures 2a–c). The intensity of PEG-SLB with CPEG = 7.5 % (Figure 2d) slowly recovered over time compare to CPEG = 0–5 %. In the PEG-SLB with CPEG = 10 %, the shape and fluorescence intensity of the photobleached region did not change with time (Figure 2e). These results indicate that increase in CPEG significantly affected the macroscopic fluidity of SLB.


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Figure 2. Fluorescence recovery after photobleaching (FRAP) processes of PEG-SLBs with CPEG of 0 – 10 %. CPEG = (a) 0 %, (b) 2.5 %, (c) 5.0 %, (d) 7.5 % and (e) 10 %. Images shown in grayscale to facilitate visualization. Quantitative FRAP measurements were performed to determine the diffusion coefficient (D) and mobile fraction (Fm) from the fluorescence recovery time course (Figure 3). The dependences of D and Fm on CPEG are summarized in Table 1. D decreased from 2.2 µm2/s to 0.74 µm2/s as CPEG increased from 0 % to 7.5 %. At CPEG = 10 %, fluorescence recovery time course could not be fitted to Eq. (1). Fm was 95 % at CPEG = 0, but decreased when CPEG > 5 %. These results indicate that the PEG-DSPE-rich domains appearing at CPEG ≥ 5% (Figure 1) had


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low fluidity and hindered lipid diffusion in SLB. Lipid diffusion was suppressed almost completely at CPEG = 10 %. The tendencies of D and Fm in this study (DOPC-SLB containing PEG-DSPE) are similar to those obtained from the FRAP analysis of in ref 25 (1-palmitoy-2dioleoyl-PC (POPC)-SLB containing PEG-1,2-dioleoyl-PE (PEG-DOPE)).

Figure 3. Fractional fluorescence recovery vs time for PEG-SLBs with various PEG-DSPE concentrations. CPEG = (□) 0 %, (△) 2.5%, (○) 5.0 %, (◇) 7.5 %, and (×) 10 %. Table 1. Diffusion coefficients (D) and mobile fractions (Fm) of SLBs containing various PEGDSPE concentrations (CPEG). CPEG [%]

D [µm2/s]

Fm

0%

2.24±0.24

0.95±0.05

2.5 %

2.19±0.19

0.93±0.03

5.0 %

1.67±0.30

0.91±0.06

7.5 %

0.74±0.39

0.64±0.11

10 %

-

0.04±0.02*

*Average fractional fluorescence intensity at 600 s after photobleaching.


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Microscopic morphology observed by atomic force microscopy The surface morphology of PEG-SLB and its dependence on CPEG were observed with AMAFM (Figures 4 and 5). The DOPC-SLB without PEG-DSPE (CPEG = 0 %) (Figure S1 of the Supporting Information) was flat and uniform except for several unruptured vesicles as described in previous studies.36,44,56 The thickness of the PEG-SLB at CPEG = 5 % was 6.7 nm (Figure S2 of the Supporting Information), which is a reasonable value of a thickness of a single lipid bilayer measured with AFM.36 At 1.0 % ≤ CPEG ≤ 5 %, there were protrusions whose density increased with CPEG (Figure 4a–d). The height and the apparent diameter (without tip size calibration) of the protrusions were 1.6 ± 0.5 nm (Figure 4f) and 50.6 ± 9.4 nm, respectively. The distance between neighboring protrusions was ~100 nm. Therefore, they were not resolved in the optical microscope images in Figure 1 (e.g. at CPEG = 2.5 %, protrusions were observed in Figure 4b, but not recognized in Figure 1b). It is reasonable to assign these protrusions to lipid clusters composed mainly of PEG-DSPE. Bulky PEG was observed higher than the surrounding DOPCrich region. The protrusions remained at the same position when the AFM scan was repeated in the same region. In SLB systems, the domains and clusters in the lipid bilayer membrane are immobile,36,37,56 although single molecules diffuse laterally. At CPEG > 5 %, the density of the protrusions decreased (Figure 4e).


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Figure 4. AFM topographies (1.0 µm × 1.0 µm) of PEG-SLBs with various PEG-DSPE concentrations. CPEG = (a) 1.0 %, (b) 2.5 %, (c) 4.0 %, (d) 5.0 % and (e) 6.0 %. (f) Cross-section profile at the white line in (c). At CPEG > 5 %, depression regions 2–3 nm in depth emerged in PEG-SLB (Figures 4e and 5), and were associated, with decrease in protrusion density. The protrusions, which were approximately 20-70 nm, were not clearly resolved with a scan size of 5×5 µm2 in Figure 5. The area fraction of this depression region increased with CPEG (Figures 5a–e). The correlation between CPEG and the area fraction of the depression regions (θdep) is plotted in Figure 6, accompanied with that of the protrusion density in Figure 4. The depression regions were not holes, but domains in PEG-SLB, because the thickness of SLB in AFM topographies is 4–6 nm generally37,45 (the assignment is discussed in detail later). The depression region had clear contrast in the phase-shift image, which is a viscoelasticity mapping (Figure S3 in the Supporting Information). These results show that the aggregation state of PEG-DSPE transformed from a


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grain-like protrusion to a two-dimensional domain and was correlated with CPEG. The depression regions shown in Figure 5 were assigned to the PEG-DSPE-rich domains presented in Figures 1c–e, on the basis of the dependence on CPEG. Both of these appeared at a threshold CPEG of ~5%. The domain had low fluidity as shown in Figure 2, and thus provided a contrast in the phase-shift image (Figure S3). Growth of the low-fluidity microdomains at CPEG > 5 % decreased macroscopic fluidity and suppressed it almost completely at CPEG = 10 % (Figures 2 and 3). The low-fluidity domains, which are, in fact, depression domains in AFM topography, segmentalized the more fluid PEG-DSPE-poor domains (Figure 5e). Isolated depression domains are recognizable in the fluorescence images (Figures 1c and 1d). In contrast, the detailed structures of the divided domains (Figure 5e) are not clearly resolved in the optical image (Figure 1f).


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Figure 5. AFM topographies (5.0 µm × 5.0 µm) of PEG-SLBs with various PEG-DSPE concentrations. CPEG = (a) 6.0 %, (b) 7.5 %, (c) 8.0 %, (d) 9.0 %, and (e) 10 %. (f) Cross-section profile at the white line in (e).

Figure 6. Dependences of the protrusion density (□) and the area fraction of the depression region (θdep) (○) on CPEG. Numerical values of the plots are summarized in Table S1 of the Supporting Information. This CPEG-dependence-based domain identification presents an apparent contradiction in that those rich in bulky PEG chains were observed as “depressions” in the AM-AFM topographies. Therefore, FM-AFM observations were made to obtain a comprehensive morphology of the PEG-rich domain.52,53 This method controls the applied force precisely in the order of pN. FMAFM (Figure 7a) and AM-AFM (Figure 7b) topographies were obtained at the same position of PEG-SLB with CPEG = 10 %. Domains higher than the surrounding region were observed in the FM-AFM image produced using an applied force of 0 pN (Figure 7a). In contrast, depressions were seen at these positions in the AM-AFM image using an applied force of 200 pN. The bulky PEG-DSPE-rich domain provided a topography similar to its morphology in FM-AFM


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observation, but caused an inverted force response in AM-AFM. In Figure 7b, the PEG-poor regions confined in the PEG-rich depression domain had a circular shape, as observed in Figure 5 (see Figure S4 in the Supporting Information).

Figure 7. AFM topographies (1.0 µm × 1.0 µm) and cross-section profiles of PEG-SLB with CPEG = 10 % obtained with (a) frequency-modulation (FM) mode at 0 pN and (b) amplitude-modulation (AM) mode at 200 pN. Both topographies were obtained at the same position. Dependence of the apparent topography on the applied force was tested by switching between 0 pN and 200 pN during FM-AFM imaging (Figure 8). The FM-AFM image at 0 pN presented with protrusions similar to those shown in Figure 7 (Figure 8a). The same area was rescanned in FM mode at 0 pN followed by an increase to 200 pN at a part of the scanning area (Figure 8b). The PEG-DSPE-rich domain appeared as a protrusion at 0 pN and a depression at 200 pN. Figures 7 and 8 show that the apparent topography of the PEG-DSPE-rich domain is significantly dependent on the applied force. These results revealed that the domains abundant in bulky PEG chains were observed as depressions in the AM-AFM topographies (Figure 5).


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Figure 8. AFM topographies (500 nm × 500 nm) of PEG-SLB with CPEG = 10 % obtained with FM mode. (a) The view field was scanned at 0 pN. (b) The applied force was changed from 0 pN to 200 pN on a portion of a PEG-DSPE-rich domain during a scan from the top to the bottom. Both topographies were obtained at the same position.

Discussion Assignment of microscopic structures observed with atomic force microscope Both fluorescence images and AFM topographies showed that PEG-DSPE-rich domains formed when CPEG > 5 % (Figures 1 and 5). AFM observation also indicated that PEG-DSPE aggregation appeared before the PEG-DSPE-rich domain formed (Figure 4). Previous studies reported that the PEG chain has two states depending on the surface density, the mushroom and brush states at lower and higher PEG density, respectively.57–61 In this study, the protrusion (Figure 4) and the depression domain (Figure 5) were assigned to the mushroom and brush states, respectively. The structure and mechanical properties of lipid bilayers containing PEGmodified lipids have been studied by QCM, neutron scattering, fluorescence microscopy and AFM force measurement.24–31 Together, these investigations defined the averaged information of macroscopic properties of PEG-SLB. The present study demonstrated the two-dimensional


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assembly of the PEG-modified lipid in the SLB system. In this study, no result implying a specific interaction between the PEG chain and substrate was obtained. D of PEG-SLB at CPEG = 0 − 5 % are ~ 2 μm2/s, which is similar to that of DOPC-SLB without PEG-lipid.44,47,51 PEGDSPE is considered existing in both leaflets of SLB as in the previous study of POPC-SLB containing PEG-DOPS.25 Decupling between the upper and lower leaflets was not observed in FRAP62 or AFM63 observations. The aggregation of the PEG-modified lipid was induced mainly by the interaction between the hydrophilic PEG chains rather those between the hydrophobic tails, because hydrophobic region was fluid in the present experimental circumstance. To validate this model, rather than using PEG-DSPE, which have saturated C18-acyl chains, other PEG-SLBs were analyzed including 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)-2000] (PEGDOPE, which have unsaturated C18-acyl chains). The results were very similar to those obtained for PEG-DSPE. FRAP measurement indicated that PEG-SLB fluidity decreased with CPEG-DOPE and depression domains were observed in AM-AFM topography at CPEG-DOPE = 10 % (Figures S5 and S6 of the Supporting Information). According to the phase diagram of DOPC and DSPE (without PEG) and assuming ideal mixing, acyl-chain-driven domain formation has rarely occurred. The ratios of the DSPE-rich gel phase at 25 °C are 0 % and 2.5 % at the DSPE concentrations of 6 mol% and 10 mol%, respectively (Figure S7 of the Supporting Information). However, PEG-DSPE-rich domains exited at CPEG = 6 % and its area fraction (θdep) was 22 % at CPEG = 10 % (Figure 6 and Table S1). Therefore, domain formation was regulated by the hydrophilic-polymer part of the PEG-modified lipid. Relationship between microscopic domain distribution and macroscopic membrane fluidity


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The macroscopic fluidity of SLB (Figures 2 and 3) was affected by the microscopic heterogeneity of morphology dependent on the PEG-DSPE concentration. The latter, in turn, was equivalent to the concentration of lipid hydrophilic polymer headgroup. The dependence of the fluidity on the PEG-DSPE concentration (Figure 3) correlates with that of the depression domain area fractions (θdep) rather than the protrusion density (Figure 6). Therefore, the PEG-DSPE-rich domains had low fluidity. These are the brighter regions in the CF-PEG-stained fluorescence image (Figure 1d) and appear as depression domains in the AM-AFM topography (Figure 5). The interaction between the PEG chains overwhelmed the thermal motion of the lipid molecules in the SLB and decreased the fluidity. The fluid fractions (Fm) in the FRAP data decreased when CPEG ≥ 7.5 % (Table 1). Nevertheless, most of the SLB remained in the relatively fluid PEGDSPE-poor region (θdep = 1.9 % at CPEG = 7.5 %, and θdep = 22 % at CPEG = 10 %; Figure 6). The PEG-DSPE-rich domains were connected to each other and divided the fluid regions (Figures 5d and 5e). Microscopic aggregation of the hydrophilic polymers lowered the macroscopic membrane fluidity in the SLB. Previous studies of block copolymer membrane systems also showed that PEG-containing amphiphilic polymers hinders the membrane fluidity.64,65 Apparent morphology of the PEG domains in atomic force microscope images The depression domains observed in the AM-AFM topography (Figure 5) should correspond to the PEG-rich regions. Nevertheless, the bulky PEG-DSPE-rich regions were lower than the periphery. The discrepancy between the AM-AFM and FM-AFM measurements (Figure 7) indicates that the apparent topography is strongly affected by the applied force. An AFM topography is the profile of a constant force between a sample and a cantilever. Therefore, AFM topographies do not always align with the geometric structure in the presence of an additional interaction like those between the hydrophilic and hydrophobic moieties in an aqueous


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solution.66–68 Hydrophilic polymers including PEG cause thermal fluctuation repulsions whose intensities depends on polymer density.69 Under AM-AFM, the PEG-rich domains were lower than the surrounding regions containing relatively less PEG-DSPE (Figure 1d). Molecular diffusion was restricted in the PEG-rich domains whereas the outer region remained fluidity. When thermal fluctuation repulsion is weaker in the PEG-rich domain than the surrounding region, at a certain applied force, the cantilever tip approaches the sample. In this applied force range, the PEG-rich domain is observed as depression. At zero applied force, FM-AFM avoids the influence of the artificial force and enables the observation of the topography (Figures 7 and 8). Kaufmann et al. plotted an AFM force curve for PEG-SLB containing DOPE-PEG.25 They reported that force-distance curve on PEG-SLB depends on the DOPE-PEG concentration. At the applied force range of 1–5 nN, which is a typical force range of AM-AFM, the tip-sample distance at 10 mol% DOPE-PEG is smaller than that measured at ≤ 8 mol % DOPE-PEG. In contrast, at applied forces < 0.1 nN, the tip-sample distance at 10 mol% DOPE-PEG is larger than that measured at ≤ 8 mol % DOPE-PEG. These results are consistent with those obtained for the AM- and FM-AFM topographies in the present study. The PEG-DSPE domains were observed as depressions under AM-AFM but as protrusions under FM-AFM. Similar force curve tendencies, shorter tip-sample distances at higher polymer-modified lipid concentrations, were reported in a lipopolysaccharide study.70 Mao et al. reported that in conventional tapping-mode (thus AM-) AFM images, the region with ~10 times higher ganglioside GM1concentration was1.5–2.6 nm lower than the surrounding region with less ganglioside GM1.18 Therefore, the interpretation of the AFM topographies in the present study are widely applicable to the domains and clusters of many lipids with hydrophilic polymers such as glycolipids. The apparent


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topography also depends on the state of the hydrophilic polymer, as well as the applied force. The other aggregation state of PEG-DSPE, which mainly appeared at CPEG ≤ 5 %, was observed as a protrusion in the AM-AFM topographies (Figure 4).

Conclusion We investigated the morphology and physical properties of PEG-SLB using fluorescence microscopes and AFM. Interactions between the PEG chains formed domains rich in PEG-DSPE wherein thermal lipid diffusion was hindered. PEG chain fluctuation in these domains created an artificial topography in the AM-AFM images. Domains rich in bulky PEG chains were observed as depressions. FM-AFM observation generated a topography reflecting the actual geometry. The discrepancies between the AM- and FM-AFM images are explained by the applied forcedependence of the thermal fluctuation repulsion. The macroscopic fluidity of the PEG-SLB is explained by the morphology and the physical properties of the PEG-DSPE-rich microdomains. These results showed that it is the hydrophilic polymer chains of lipid head groups, rather than their hydrophobic tails, that play a dominant role in determining membrane fluidity and microscopic morphology of the PEG-SLB. These hydrophilic-polymer-derived interactions may determine the roles of glycolipids and lipopolysaccharides in the function of the cell membranes of living cells and bacteria. The methodology and the interpretation in the present study support the investigation of these lipids.

ASSOCIATED CONTENT


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Supporting Information. The Supporting Information is available free of charge on the ACS Publications website at DOI: *************** Fluorescence images and AFM topographies of SLB containing PEG-DOPE, a table summarizing the numerical values of Figure 6, calculated phase diagram of ideally mixed DOPC+DSPE-bilayer (PDF)

AUTHOR INFORMATION Corresponding Author *E-mail: [email protected]. Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT This work was supported by JSPS KAKENHI Grant Numbers JP15H03768 and JP15H00893, JST-CREST Grant Number JPMJCR14F3, JST-A-STEP, EIIRIS Project from TUT, and CHOZEN Project, Kanazawa University.


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Figure 1. Fluorescence images of the PEG-SLBs with CPEG = 0–10 %. (a, b) PEG-SLB with (a) CPEG = 0 % (DOPC-SLB) and (b) CPEG = 2.5 % labeled with Rb-DPPE (Ex/Em: 560/583 nm), (c, d) PEG-SLB with CPEG = 5 % labeled with Rb-DPPE and CF-PEG-DSPE (Ex/Em: 485/523 nm), (c) Rb-DPPE and (d) CF-PEG-DSPE images obtained at the same position. (e) Merged image of (c) and (d). Insert: magnified image of the dotted square region. Representative PEG-DSPE-rich green regions are indicated with a white arrow. Dark defect is indicated by a black arrow. (f) PEG-SLB with CPEG = 10 % labeled with Rb-DPPE. 969x477mm (96 x 96 DPI)

ACS Paragon Plus Environment

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Figure 2. Fluorescence recovery after photobleaching (FRAP) processes of PEG-SLBs with CPEG of 0–10 %. CPEG = (a) 0 %, (b) 2.5 %, (c) 5.0 %, (d) 7.5 % and (e) 10 %. Images shown in grayscale to facilitate visualization. 774x867mm (96 x 96 DPI)


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Figure 3. Fractional fluorescence recovery vs time for PEG-SLBs with various PEG-DSPE concentrations. CPEG = (□) 0 %, (△) 2.5%, (○) 5.0 %, (◇) 7.5 %, and (×) 10 %. 500x453mm (96 x 96 DPI)


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Figure 4. AFM topographies (1.0 µm×1.0 µm) of PEG-SLBs with various PEG-DSPE concentration. CPEG = (a) 1.0 %, (b) 2.5 %, (c) 4.0 %, (d) 5.0 % and (e) 6.0 %. (f) Cross-section profile at the white line in (c). 946x581mm (96 x 96 DPI)


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Figure 5. AFM topographies (5.0 µm×5.0 µm) of PEG-SLBs with various PEG-DSPE concentrations. CPEG = (a) 6.0 %, (b) 7.5 %, (c) 8.0 %, (d) 9.0 %, and (e) 10 %. (f) Cross-section profile at the white line in (e). 948x561mm (96 x 96 DPI)


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Figure 6. Dependences of the protrusion density (□) and the area fraction of the depression region (θdep) (○) on CPEG. Numerical values of the plots are summarized in Table S1 of the Supporting Information. 601x478mm (96 x 96 DPI)


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Figure 7. AFM topographies (1.0 µm × 1.0 µm) and cross-section profiles of PEG-SLB with CPEG = 10 % obtained with (a) frequency-modulation (FM) mode at 0 pN and (b) amplitude-modulation (AM) mode at 200 pN. Both topographies were obtained at the same position. 528x430mm (96 x 96 DPI)


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Figure 8. AFM topographies (500 nm × 500 nm) of PEG-SLB with CPEG = 10 % obtained with FM mode. (a) The view field was scanned at 0 pN. (b) The applied force was changed from 0 pN to 200 pN on a portion of a PEG-DSPE-rich domain. Both topographies were obtained at the same position. 516x285mm (96 x 96 DPI)


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Morphology and Physical Properties of Hydrophilic-Polymer-Modified

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