Imaging Characterization of Cluster-Induced Morphological Changes

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Imaging Characterization of Cluster-Induced Morphological Changes of a Model Cell Membrane Hideki Nabika, Aya Sakamoto, Toshinori Motegi, Ryugo Tero, Daiki Yamaguchi, and Kei Unoura J. Phys. Chem. C, Just Accepted Manuscript • DOI: 10.1021/acs.jpcc.5b08014 • Publication Date (Web): 09 Feb 2016 Downloaded from http://pubs.acs.org on February 17, 2016

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The Journal of Physical Chemistry

Imaging Characterization of Cluster-Induced Morphological Changes of a Model Cell Membrane Hideki Nabika1*, Aya Sakamoto1, Toshinori Motegi2, Ryugo Tero2, Daiki Yamaguchi1, and Kei Unoura1

1

Department of Material and Biological Chemistry, Faculty of Science, Yamagata

University, 1-4-12 Kojirakawa, Yamagata 990-8560, Japan 2

The Electronics-Inspired Interdisciplinary Research Institute, Toyohashi University of

Technology, 1-1 Hibarigaoka, Tempaku-cho, Toyohashi, 441-8580, Japan

*e-mail: [email protected]

Keywords: antimicrobial material, polyoxometalate, cluster, membrane, lipid bilayer

1

ACS Paragon Plus Environment


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Abstract Understanding the activity of nanomaterials at the lipid bilayer surface can provide key information for the feasible designing of functional bioactive agents. Herein, we used micro- and nanoscopic imaging techniques to evaluate the activity of nanometer-sized inorganic clusters, and report that destruction of the lipid membrane is induced by a cluster-induced morphological change on the membrane surface. As model experiments, we used the Keggin-type polyoxometalate (POM) SiW12O404- for the inorganic cluster, and

a

1,2-dimyristoyl-sn-glycerol-3-phosphatidylcholine

(DMPC)

and

egg

phosphatidylcholine (EPC) bilayer for the cell membrane. Imaging experiments revealed vigorous desorption of the lipid bilayer from solid substrate by the formation of POM–lipid assembly through a supramolecular-type assembly process in which electrostatic and hydrophobic interactions between the POM and lipid determines the efficiency and dynamics of assembly formation and thereby determines lipid desorption. Furthermore, maximum efficiency of lipid desorption was found at the phase transition temperature. This phase dependency was explained by the formation of a “leaky interface” between the gel and fluid domains, in which freedom in the conformational change of lipids during the formation of the POM–lipid assemblies becomes maximal.

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Introduction The cell membrane is an important field for biological reactions. Physical insights into reactions on the cell membrane are necessary to design biologically active materials.1,2 Anti-microbial peptides (AMP), which play roles in wound healing and modulation of the innate and adaptive immune systems, are among the typical biological materials exhibiting characteristic interactions on the cell membrane. AMPs demonstrate their antimicrobial activity by interacting directly with the cell membrane. One well-established mechanism of the antimicrobial activity of AMPs is via pore formation on the cell membrane,3 which disrupts its important role as a barrier between the cell interior and the extracellular milieu. Materials showing characteristic interactions with the cell membrane have been explored by many researchers. In this regard, recent efforts have been devoted to exploit the activity of nanoparticles on biological membranes4-6 with particular focus on metal,7-12 metal oxide,13-18 semiconductor,19-21 and polymeric nanoparticles22-25 because these nanoparticles are often stable and synthesizable using relatively low-cost processes. Depending on the material, size, and surface functionality, diverse interactions of nanoparticles with membranes have been reported. For example, an AMP-like ability characterized by a leakage phenomenon was reported when positively charged gold and titanium oxide nanoparticles were mixed with liposomes.8 The interaction between nanoparticles and membranes can be tuned via the flexibility of a surface molecule.26 Moreover, the adsorption of polymeric nanoparticles onto the membrane surface induces the reconstruction of lipid molecules at the point of nanoparticle adsorbtion.22 This reconstruction then alters the thermodynamic stability and disturbs the phase state. Collectively, these reports have revealed the ability of 3



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nanomaterials to interact with and denature cells. Furthermore, recent developments in synthesis methodologies have enabled the fabrication of nanoparticles with a wide variety of sizes, shapes, charges, and surface functionalities, paving the way for the development of novel antimicrobial nanoparticles with unique and diverse properties. However, even nanoparticles developed by well-controlled synthesis will occasionally have a non-zero size distribution.27,28 Such variation in size or shape may result in kinetic or thermodynamic heterogeneity with respect to the activity toward the cell membrane.29,30 The recent development of nanometer-sized inorganic cluster materials could help overcome these extensive size and shape distributions, as these materials are characterized by single and definitive compositions and structures. Gold clusters have been among the most vigorously investigated clusters to date,31 but the synthesis, purification, and separation of sufficient amounts of a single component to gain insights into the chemical, physical, and biological natures of each cluster remain challenging.32 Thus, one of the most potent next-generation antimicrobial material candidates is a metal-oxide cluster that would enable large-scale production at the kilogram scale. Two independent research groups have succeeded in revealing the membrane-targeting ability of one such metal-oxide cluster, polyoxometalate (POM), by using liposomes as model cell membranes.33,34 The adsorption of POM onto a lipid bilayer membrane induces a morphological change of the membrane, which depends on the type of POM used. In particular, when a Keggin-type POM is used, the membrane is gradually broken down, and most of the liposomes collapse. The mechanism of such highly destructive activity has been proposed to occur in the following three steps (Figure 1): (i) adsorption of POM onto the membrane surface, (ii) reorientation of a lipid molecule 4


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to form lipid–POM assemblies on the membrane, and (iii) desorption of the assemblies into the bulk water phase.33 Despite POM’s promising membrane-targeting ability, direct evidence of the proposed mechanism based on the formation and desorption of POM–lipid assemblies is lacking. In the present study, we directly observed the morphological changes in a lipid bilayer membrane upon POM addition based on fluorescence and atomic force microscopy (AFM) for micro- and nanoscale morphological characterization, respectively. In addition, the phase-dependent membrane-targeting ability of POM was investigated to explore its membrane selectivity, i.e., cell selectivity, which could further enhance the applicability of POM as a novel bioactive material.

Figure 1. Schematic illustration of the proposed destructive activity of polyoxometalate (POM) towards the cell membrane. POM and lipid molecules are shown in yellow and gray, respectively. Adsorption of POM on the membrane surface leads to the formation of a POM–lipid assembly, which finally desorbs from the membrane surface.

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Materials and Methods Materials. A commercially available Keggin-type POM, H4[SiW12O40] (Wako Pure Chemical

Industries

Ltd.),

was

used

without

further

purifications.

1,2-Dimyristoyl-sn-glycerol-3-phosphatidylcholine

(DMPC),

1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine-N-(7-nitro-2-1,3-benzoxadiazol-4-y l) (NBD-PE), and egg phosphatidylcholine (EPC) were purchased form Avanti Polar Lipids (Alabaster, AL, USA) and used as received. The NBD-PE/DMPC bilayer and EPC bilayer were formed on a clean substrate for observations with a fluorescence microscope

(BX-53,

Olympus

Co.,

Ltd.,

Japan)

equipped

with

a

temperature-controllable stage (TP-CHS-B13V, Tokai Hit Co., Ltd., Japan) and by an atomic force microscope (Agilent Technologies, Inc., formerly Molecular Imaging, Corp.) equipped with a closed-loop scanner, respectively. Morphological changes in the NBD-PE/DMPC and EPC bilayers resulting from the addition of an appropriate amount of a Tris/HCl (pH 7.4)-buffered H4[SiW12O40] solution were monitored.

Fluorescence microscope observations. For fluorescence microscope observations, the NBD-PE/DMPC bilayer was used. A small amount of NBD-PE/DMPC-chloroform solution was placed onto a clean glass substrate. After removing the chloroform by drying, the substrate cast with the NBD-PE/DMPC was immersed in a petri dish containing 150 mM Tris/HCl (pH 7.4) buffer and incubated at 40°C overnight. The petri dish was then transferred to a temperature-controllable stage (TP-CHS-B13V) mounted 6


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on a fluorescence microscope (BX-53). An appropriate amount of POM buffer solution was added to the petri dish at desired concentrations. The fluorescence images were acquired by a CCD camera (DP-21, Olympus Co., Ltd., Japan).

AFM observations. EPC was used for conducting AFM observations at room temperature. Supported lipid bilayers (SLBs) were formed on the substrate surfaces by the vesicle fusion method.35,36 A chloroform solution of EPC (10 mg/ml) and an ethanol solution of BODIPY-H-PC (4.6 × 10-2 mg/ml) were mixed in a glass vial at the required amount and ratio, and the solvents were evaporated by N2 blow-down followed by evacuation in a vacuum desiccator for over 6 h. We prepared a multilamellar vesicle suspension with a 0.4 mM lipid concentration by agitating the vacuum-dried lipid films in a buffer solution (150 mM Tris/HCl buffer, pH 7.4). The suspension was repeatedly frozen in liquid nitrogen and thawed in a 45°C water bath five times and sequentially extruded through an 800-nm polycarbonate filter (Whatman Inc.) five times and through a 100-nm polycarbonate filter six times to obtain unilamellar vesicles. The extruded vesicle suspension was diluted to 0.1 mM and sonicated in a water-bath ultrasonic cleaner to produce smaller vesicles. The thermally oxidized SiO2/Si substrates were immersed in the sonicated vesicle suspension and incubated at 45°C for 60 min. After the incubation, the excess vesicles were washed out in the liquid phase by exchanging the suspension with the buffer solution. AFM topographies were obtained using PicoScan2500 (Agilent Technologies Inc., formerly Molecular Imaging Corp.) equipped with a closed-loop scanner. SLB observation was performed in the buffer solution in acoustic-AC

mode

(conventional

tapping

mode)

7


with

a

microcantilever


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(OMCL-AC240TN-C3, Olympus). All images were obtained with 256 scan points, 256 scan lines, and a scan rate of 0.2 Hz in the buffer solution at room temperature. AFM image processing was performed with the commercial software package SPIP 3.0 (Image Metrology).

Results and Discussion Fluorescence microscope observations. Fluorescence microscope observations were conducted for the NBD-PE/DMPC bilayer at the phase transition temperature (24.5°C ± 0.5°C) because lipid desorption was most apparent under these temperatures, as discussed below. First, we investigated the morphological changes in the NBD-PE/DMPC bilayer induced by the addition of POM at different concentrations. Control experiments verified that any morphological changes were not observed by pH change or tungstate addition that would be induced by POM addition (Supporting Information Figure S1). When the final POM concentration was below 38.4 µM, no change in fluorescence intensity other than photobleaching, and thus, morphology, was observed before (Figure 2A) or after (Figure 2B) POM addition. It should be noted that photobleaching was observed during the observation without the addition of POM, i.e., the 0 µM condition, which is discussed in detail below. The large-area fluorescence intensity histogram indicated that the bilayer exhibited a fluorescence intensity maximum at around 50 counts, whereas a small peak appearing at 15 counts represented the background intensity. Thus, the net fluorescent intensity of the intact NBD-PE/DMPC bilayer was 35 counts. The addition of POM at 384 µM and 500 µM induced two distinct changes: a decrease in fluorescence intensity of the whole area, and 8


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the appearance of bright patches a few micrometers in diameter on the dark surrounding

Figure 2. Fluorescence microscope images and fluorescence intensity analysis. Fluorescence

microscope

images

of

the

1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine-N-(7-nitro-2-1,3-benzoxadiazol-4-y l) (NBD-PE)/1,2-dimyristoyl-sn-glycerol-3-phosphatidylcholine (DMPC) bilayer on a glass substrate (A) before and (B) 10 min after the addition of polyoxometalate (POM) solutions at the indicated concentrations. (C) Fluorescence intensity histograms (black) before and (red) after the POM addition. (D) Fluorescence intensity histograms showing the intensity range between 40 and 80.

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region with reduced intensity. These changes were noted within a few minutes after POM addition (see Supporting Information Figure S2); this induction period of a few minutes might correspond to the time required for POM to diffuse to the surface of the bilayer in the petri dish, because the solution was not stirred during the observation period. A further increase in POM concentration to 3840 µM resulted in efficient removal of the lipids from the substrate, as indicated by the reduction in fluorescence intensity down to near-background levels of the whole area.

10


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Figure 3. (A) Successive fluorescence microscope images and (B) intensity line profiles for the white dashed line in the fluorescence microscopy images of the NBD-PE/DMPC bilayer on a glass substrate taken every 5 min in a 384 µM polyoxometalate (POM) solution at 24.4°C. The contrast of each image was tuned for clarity.

The observed decrease in the fluorescence intensity implies desorption of lipid molecules from the bilayer deposited on the glass substrate. It should be noted that the POM used in the present study does not exhibit significant fluorescence-quenching activity toward NBD dye molecules, which was confirmed by mixing POM and 4-fluoro-7-nitrobenzofurazan (NBD-F) in ethanol (see Supporting Information Figure S3). Intensity reduction in the bilayer can be seen more clearly by focusing on the bilayer edge region (Figure 3) because the reaction may proceed from the bilayer edge, similar to the hydrolysis of lipids by phospholipase.37 As expected, lipid desorption was clearly observed to be triggered at the bilayer edge, leaving small bilayer domains. The fluorescence intensity decreased down to near-background level, in which the presence of small bilayer domains that could not be resolved by fluorescence microscopy yielded some fluorescence at these regions. Reduction in the fluorescence intensity down to near-background level indicates that POM addition caused bilayer desorption from the glass substrate. Furthermore, pore formation was also evident at several points. At the pores, background-level fluorescence intensity was observed, indicating almost complete bilayer desorption. These observations suggest the ability of POM to desorb the lipid bilayer, similar to nanoparticle-induced bilayer desorption.38 With regard to bilayer desorption, the interaction between POM and lipids might play a crucial role. Negatively charged POM has the ability to form assemblies 11



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with surfactants that have a positive head group via an electrostatic interaction. The cationic surfactant can then replace the countercations on POM, which forms a self-assembled structure called a surfactant-encapsulated cluster (SEC).39-43 SECs are generally formed and characterized in organic solvents. The hydrophobic tails of the surfactant adsorbed on POM orient in the organic solvent. However, our system focused on the formation of POM–lipid assembly in an aqueous phase, indicating that SEC in the organic solvent cannot be stably dispersed in water due to its hydrophobic nature. Thus, it is reasonable to assume that the formation of non-hydrophobic POM–lipid assemblies occur to lower the free energy in water. Thus, one can expect an interdigitated configuration of lipid molecules.44 In particular, small materials with high curvature, e.g., nanoparticles or clusters, promote the formation of an interdigitated configuration on the particle surface. One example can be found with respect to water-soluble gold nanoparticles encapsulated by an interdigitated molecular layer consisting of lipid and alkanethiol.45 By interdigitating an amphiphilic lipid to a hydrophobic alkanethiol adsorbed on the nanoparticle surface, the nanoparticles can gain solubility by orienting the hydrophilic moiety of the hydrophobic tails at the aqueous phase. Similarly, by adopting an interdigitated configuration of lipids on the POM surface, the charged hydrophilic head group can orient at both the POM surface and water phase. This enhances the solubility and stability of POM–lipid assemblies in water. The driving force for the formation of the interdigitated lipid configuration on POM depends on the POM–lipid electrostatic interaction and the lipid–lipid hydrophobic interaction; no covalent linkage would be associated. It is also likely that the hydrophobic interaction between lipids adsorbed on different POMs can cause a linkage between these two POM–lipid assemblies. Successive linkage would form large 12


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aggregates on the lipid layer. Upon linkage among POM–lipid assemblies, the configuration may change among unstructured aggregate, micelle, and vesicles, either on the bilayer surface or in the water phase, to lower the free energy. Furthermore, since the number of lipids that can interact with POM is determined by the charge of POM, the efficiency of POM–lipid assembly and its aggregate formation, and thus, lipid desorption efficiency, are expected to be tunable by adjusting the structure and charge of POM. Thus, this kind of lipid desorption undergoes a cooperative assembly among many POMs and lipid molecules to form large aggregates, different from nanoparticle-induced desorption, which proceeds via the interaction between a single nanoparticle and bilayer envelope.38 This suggestion can also be expected from the energetics viewpoint. The free energy change ∆G upon nanoparticle-induced lipid desorption was discussed in terms of particle–bilayer adhesion energy, bilayer bending energy, and bilayer interfacial (defect) energy. On the basis of numerical analysis for ∆G, the minimum particle diameter to satisfy the condition ∆G < 0 was estimated to be a few tens of nanometers, which is much larger than the size of the POM (~1 nm) used in the present study. Thus, there would be a critical transition in nanoparticle–lipid interaction when nanoparticle size goes down to a single-nanometer scale. Instead of nanoparticle-type interaction, the energetic landscape for POM–lipid assembly process would obey a similar mechanism of supramolecular assembly, in which constituent ions and molecules interact via weak interaction and form a large assembly as one of a thermodynamically stable state.46 The electrostatic interaction between POM and lipid and the hydrophobic interaction among lipids would induce either step-by-step assembly (isodesmic model) or abrupt assembly above a critical POM concentration (cooperative model).47 As demonstrated by the results, they grow into micrometer-scale 13



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assemblies and desorb from the bilayer surface when they are sufficiently soluble into the aqueous phase, which would explain the observed POM-induced lipid desorption. Our finding offers physical insights into nanomaterial–bilayer interaction in that the mechanism has critical transition when the size of nanomaterials goes down to single nanometer region. Bright patches seen in Figure 2 (under 384 µM and 500 µM conditions) would corresponds to these large POM-lipid assembly or aggregate. At POM concentrations of 3.84 µM and 38.4 µM, a sharp cutoff was observed at a fluorescence intensity of 60 counts before and after the POM addition (Figure 2C and D). A slight shift in the cutoff intensity before and after POM addition is attributable to power fluctuations of the Hg lamp used as the excitation light source (slight power fluctuations were unavoidable with the Hg lamp used in the present experimental setup). In contrast, a clear shoulder to a higher intensity of up to 80 counts appeared at POM concentrations of 384 µM and 500 µM (depicted with red arrows in Figure 2D), which corresponds to the intensity of the bright patches. This result indicates that the bright patches do not represent a bilayer domain that was left after POM addition but rather reflect additional architectures that were newly formed on the bilayer after POM addition. Since the addition of POM was the trigger of the formation of the bright patches, they could be attributed to the formation of POM–lipid aggregates as discussed above. Since POM–lipid assemblies and aggregates are known to expand up to a few micrometers,39,40 the largest bright patches with the size of a few micrometers would correspond to such large aggregates.

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Figure 4. Temporal changes in bright patches observed for NBD-PE/DMPC in a 384 µM polyoxometalate (POM) solution. (A) Translocation and (B) morphology change and desorption of the bright patches. The contrast of each image was tuned for clarity. The scale bar is 10 µm.

Furthermore, the bright patches exhibited morphological changes over time. Some bright patches exhibited slow diffusion (Figure 4A), although this translocational movement was not observed for all the patches; several patches did not diffuse at all throughout the observation period. There is currently no criterion, e.g., size or shape, to clearly distinguish mobile and immobile patches. Furthermore, some patches gradually changed their shape and occasionally disappeared or were desorbed (Figure 4B). Morphological changes would be followed by the rearrangement of large POM–lipid assemblies. These temporal morphological changes support our suspicion that the observed bright patches represent an independent architecture formed on the underlying bilayer.

Phase dependency. From the fluorescence microscopy images, the lipid desorption 15



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ratio can be quantitatively analyzed from a decrease in the fluorescence intensity. We defined the desorption ratio as R, which was calculated from the decrease in the fluorescence intensity of the membrane before and 10 min after the addition of POM. Without POM, fluorescence microscopy observation for 10 min resulted in a non-zero R value; the fluorescence intensity was reduced by 20% at any temperature ranging from 10°C (gel phase) to 40°C (fluid phase) (Figure 5A). The observed intensity reduction was caused by photobleaching of the NBD moiety. At a POM concentration of 3.84 µM,

Figure 5. Fluorescence image analysis. (A) Temperature-dependent lipid desorption ratio

(R)

for

the

1,2-dimyristoyl-sn-glycero-3-phosphoethanolamine-N-(7-nitro-2-1,3-benzoxadiazol-4-y l) (NBD-PE)/1,2-dimyristoyl-sn-glycerol-3-phosphatidylcholine (DMPC) bilayer in polyoxometalate (POM) solutions at different concentrations. (B) Plot of the immobile fraction (IF) versus diffusion coefficient (D) for the NBD-PE/DMPC bilayer at 24°C (red) and 40°C (blue). The POM concentrations are: ▲ 0 µM, ■ 3.84 µM, and ● 500 µM.

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the R value was almost the same as that observed under the photobleaching effect, indicating that no detectable desorption was induced at any temperature. An increase in POM concentration yielded a detectable increase in R. Furthermore, temperature dependence was evident. For the 384 µM and 500 µM conditions, maximum R values of around 80% and 100% were observed near the phase-transition temperature, respectively. Both decreasing and increasing temperature from the phase-transition temperature reduced R, indicating that desorption of lipids from the membrane occurs more effectively under the phase transition state. The temperature dependency of R can be explained as being akin to the temperature dependency in the permeability of ions,48,49 small molecules,50,51 and peptides.52 It is well known that gel and fluid phases have different chemical and physical properties. The coexistence of the gel and fluid domains at the phase-transition state provides an interface between these dissimilar domains. Some ions or molecules have a maximum permeability at the phase-transition temperature,48-52 where domain interfaces are formed. A current density experiment conducted to monitor ion permeability demonstrated that the phase-transition state has a relatively long period of the “pore” state for ion transport.48 Another experiment using small molecules suggested that the acyl chain located at the domain interface has sufficiently large density fluctuation to allow for the rapid transmembrane diffusion of small molecules.50 These reports clarified the specific nature of the domain interface that has a loose packing structure and high ability to accommodate other species by changing the conformation of lipid molecules, which is sometimes referred to as a leaky interface.53 As discussed above, the destructive activity of POM is induced by the transition from a POM-on-bilayer state to a POM–lipid assembly. Freedom in the conformational change 17



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of a lipid molecule in the membrane would therefore be an important parameter to govern the efficiency of this transition process. Since a leaky interface at the phase-transition regime is expected to confer higher freedom for conformational changes during the transition into the POM–lipid assembly, the observed temperature dependency of R can be explained by the presence of a leaky interface, similar to the temperature dependency observed with respect to permeability. To further confirm the temperature-dependent activity of POM, we analyzed the diffusivity of lipid molecules after the addition of POM via the fluorescence recovery after photobleaching (FRAP) technique (see Supporting Information Figure S4). FRAP provides two independent parameters: the diffusion coefficient (D) and the immobile fraction (IF). At 40°C, the D value of the fluidic DMPC bilayer decreased from 5.7 to 2.0 µm2/s when the POM concentration increased from 3.84 to 500 µM (Figure 5B). Since the D of DMPC at around 40°C has been reported to be 6–9 µm2/s,54,55 the observed value of 5.7 µm2/s at a POM concentration of 3.84 µM corresponds to a nearly intact state of the DMPC bilayer, which is consistent with the results for R shown in Figure 5A. The decrease in D with increasing POM concentration seems to result from the formation of defect sites due to bilayer desorption, as discussed above. Lipid desorption disrupts the continuity of the lipid membrane, which acts as a diffusion barrier for lipids. D is known to decrease when the observation period is sufficiently long to allow the lipid to encounter defective sites during their diffusion,54,55 which has been used as a measure to evaluate a compartment formed by the membrane skeleton network.56 In our experiment, the FRAP observation period was 25 min, and each lipid diffused to around 3000 µm2 (i.e., 50 × 50 µm2) at 2.0 µm2/s. Thus, the decrease in D at the high POM concentration condition indicates the presence of at least 18


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one defective site per 50 × 50-µm2 area. This estimation is reasonable considering the results of fluorescence microscopy observations, in which lipids were removed at many sites in nearly 50 × 50-µm2 areas of 500 µM. On the other hand, IF was almost unchanged under the experimental conditions at 40°C. When the experiments were conducted at the phase-transition temperature, at the lowest POM concentration, D was about 2.9 µm2/s, which is comparable to previously reported values.57,58 Increasing the POM concentration resulted in a decrease in D to 2.2 µm2/s, which is similar to the results observed at 40°C. A further increase to 500 µM POM resulted in a slight decrease in D and a significant increase in IF. Appearance of a non-zero IF value indicates intense growth of bilayer defects and the formation of an isolated bilayer domain that cannot undergo diffusive lipid exchange with its surroundings;59 this domain could be formed by the growth of defective sites in both size and number, so that they ultimately fuse into a network-like defect. Thus, the FRAP experiments further confirmed that temperature, i.e., phase condition, is an important parameter for POM to have a destructive interaction with lipid membranes.

AFM observations. We also conducted AFM observations of the bilayers after the interaction with POM. AFM must be performed under room temperature, and DMPC at room temperature (i.e., at the phase transition temperature) was found to be unstable and fragile in the presence of POM. Thus, we could not exclude the potential of unfavorable artifacts caused by physical stress during the tip scanning process. To this end, we used an EPC bilayer, which has a phase transition temperature far below room temperature, so that the bilayer could be maintained in fluidic phase throughout the observations. At the lowest POM concentration of 38.4 µM, we could not observe any 19



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change in the AFM image, consistent with the fluorescence microscopy observations (see Supporting Information Figure S5). The increase in POM concentration to 384 µM caused the formation of defect-like pores. However, as discussed later, the EPC bilayer is more stable against POM addition, and lipid desorption was not found to be significant compared with DMPC by fluorescence microscopy. However, the change became evident when the POM concentration increased to 3840 µM, wherein lipids were vigorously desorbed and tiny domains were left on the substrate (Figure 6A). Since many of these domains were smaller than 1 µm, they could not be detected under the fluorescence microscope at such a high POM concentration. Specifically, the AFM images revealed the presence of domains 6–7 nm in height and 0.1–1 µm in diameter

Figure 6. Atomic force microscopy (AFM) observations. (A) AFM image of the egg phosphatidylcholine (EPC) bilayer in a 3840 µM polyoxometalate (POM) solution. (B) The height profile for the region is shown with a white dashed line in (A).

(Figure 6B), representing the bilayer domains that remained after POM addition. The AFM image and height line profile also exhibited the presence of a trace amount of tiny 20


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domains 2–3 nm in height, corresponding to a layer of residual lipid molecules. Thus, the dominant component left on the substrate after exposure to the 3840 µM POM solution were small EPC bilayer domains measuring less than 1 µm in diameter. Furthermore, bright (i.e., larger height) objects were frequently found on small bilayer domains. These bright objects likely correspond to small remnant POM–lipid assemblies that were maintained even under the high POM concentration condition, with a height of 5–20 nm (see Supporting Information Figure S6). By combining the information obtained from the fluorescence microscopy and AFM observations, the process of POM-induced morphological change is proposed as follows. Once POM is added into the petri dish, it diffuses to the surface of the bilayer membrane, where an electrostatic interaction causes the adsorption of POM. The adsorbed POM forms POM–lipid assemblies according to the isodesmic-type supramolecular assembly process. Most of the POM–lipid assemblies desorb, and only small bilayer domains are left on the substrate. At the same time, some of the initially formed POM–lipid assemblies can remain on the membrane surface, where they undergo growth, shrinkage, and diffusion. At especially high POM concentrations, just tiny bilayer domains holding nanometer-scale POM-lipid assemblies are left; most lipids are desorbed from solid substrate surface. It should also be noted that the fluorescence intensity histogram of DMPC with 3840 µM POM exhibited almost no fluorescence, suggesting that no lipids were left on the substrate (Figure 2). On the other hand, in the case of EPC with 3840 µM POM (Figure 6), about 17% (the lipid area occupied 65,000 pixels of a total area of 380,000 pixels) of the substrate surface was still covered with a bilayer domain, which should result in detectable fluorescence intensity if the same amount of DMPC domains were 21



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left on the substrate. This inconsistency can partly be explained by considering the phase dependency of the reaction. At room temperature, DMPC and EPC are in the phase transition and fluid states, respectively; thus, it is reasonable to expect higher destructive activity on DMPC than EPC for the same POM concentration. Such composition-dependent efficiency shows good potential for the cell selectivity of cluster-induced antimicrobial functionality.

Conclusion POM clusters can induce morphological changes in a POM–lipid assembly according to the supramolecular-type assembling process. Electrostatic interaction and hydrophobic interaction among POM and lipid would induce successive assembly of these constituents, which can grow into micrometer-scale assembly. Then, they desorb from the bilayer surface when they are sufficiently soluble into the aqueous phase, which would be the origin of POM-induced lipid desorption. Since a weak interaction between POM and lipid governs the assembly process, POM can be suitably designed to control its activity at the cell interface. The maximum lipid-desorption efficiency was observed at the main phase transition temperature of the bilayer, indicating that a presence of gel/fluid interface with a loose packing structure plays an important role for POM-induced lipid desorption. This phenomenon was explained by the similarity with the temperature dependency of drug permeability through the lipid bilayer.

Acknowledgments. This work was supported by Grant-in-Aid for Challenging Exploratory Research 26600021 and Nippon Sheet Glass Foundation for Materials Science and Engineering 22


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(NSG Foundation).

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Table of Contents Graphic

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Imaging Characterization of Cluster-Induced Morphological Changes

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