Atomic force microscopy study on stiffness of nanosized liposomes

Subscriber access provided by Kaohsiung Medical University

Interface Components: Nanoparticles, Colloids, Emulsions, Surfactants, Proteins, Polymers

Atomic force microscopy study on stiffness of nanosized liposomes containing charged lipids Yuki Takechi-Haraya, Yukihiro Goda, and Kumiko Sakai-Kato Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b01121 • Publication Date (Web): 05 Jun 2018 Downloaded from http://pubs.acs.org on June 5, 2018

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

Atomic force microscopy study on stiffness of nanosized liposomes containing charged lipids Yuki Takechi-Haraya, Yukihiro Goda, and Kumiko Sakai-Kato* National Institute of Health Sciences, 3-25-26 Tonomachi, Kawasaki-ku, Kawasaki City, Kanagawa 210-9501, Japan *To whom correspondence should be addressed: Dr. Kumiko Sakai-Kato, Ph.D. Division of Drugs, National Institute of Health Sciences 3-25-26 Tonomachi, Kawasaki-ku, Kawasaki City, Kanagawa 210-9501, Japan Tel: +81-44-270-6510 Fax: +81-44-270-6510 E-mail: [email protected]

ACS Paragon Plus Environment

1

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 2 of 34

ABSTRACT

It has recently been recognized that the mechanical properties of lipid nanoparticles play an important role during in vitro and in vivo behaviors such as cellular uptake, blood circulation, and biodistribution. However, there have been no quantitative investigations of the effect of commonly-used charged lipids on the stiffness of nanosized liposomes. In this study, by means of atomic force microscopy (AFM), we quantified the stiffness of nanosized liposomes composed of neutrally charged lipids combined with positively or negatively charged lipids while simultaneously imaging the liposomes in aqueous medium. Our results showed that charged lipids, whether negatively or positively charged, have the effect of reducing the stiffness of nanosized liposomes, independently of the saturation degree of the lipid acyl chains; the measured stiffness values of liposomes containing charged lipids are 30–60% lower than those of their neutral counterpart liposomes. In addition, we demonstrated that the Laurdan generalized polarization (GP) values, which are related to the hydration degree of the liposomal membrane interface and often used as a qualitative indicator of liposomal membrane stiffness, do not directly correlate with the physical stiffness values of the liposomes prepared in this study. However, our results indicate that direct quantitative AFM measurement is a valuable method to gain molecular-scale information about how the hydration degree of liposomal interfaces reflects (or does not reflect) liposome stiffness as a macroscopic property. Our AFM method will contribute to the quantitative characterization of the nano-bio interaction of nanoparticles, and to the optimization of the lipid composition of liposomes for clinical use.

Keywords: stiffness; nanosized liposome; atomic force microscopy; charged lipid; hydration degree


2

Page 3 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

INTRODUCTION Liposomal formulations are a promising strategy for drug delivery to advance the treatment of various diseases.1 Liposomes are spherical lipid-bilayer vesicles typically composed of amphiphilic, neutrally charged lipids, such as phosphatidylcholines. They assemble mainly via hydrophobic attraction in aqueous solution, and are used as drug carriers because of their ability to encapsulate drug molecules in the vesicular structure.2 In particular, nanosized liposomes with hydrodynamic diameter (or size) of around 100 nm are known to exhibit well-dispersed stability and efficient accumulation into tumor tissue. These properties stimulate active research and development of liposomal drug products for cancer therapy.1,2 Many reports have shown the advantages of using positively or negatively charged lipids in addition to neutrally charged lipids for liposomal formulations.3–13 For example, in the formulation of AmbisomeⓇ, well-dispersed, stable liposomes with the ability to target fungi have been achieved by use of the negatively charged phospholipid 1,2-distearoyl-sn-glycero-3phospho-(1′-rac-glycerol) (DSPG), thereby improving the therapeutic efficacy and reducing the toxicity.3

Elsewhere,

liposomes

containing

cationic

lipids

such

as

1,2-dioleoyl-3-

trimethylammoniumpropane (DOTAP) have been identified as useful carriers of drugs, including not only low-molecular-weight chemicals but also nucleotides such as DNA and siRNA, which are unstable in the blood.6–11 For the successful development of liposomal formulations for clinical use, it is crucial to devise appropriate lipid compositions to achieve the desired properties, such as drug release rate, cellular uptake efficiency, circulation time, and biodistribution.14 Therefore, the establishment of methodologies to characterize liposomes with various lipid compositions and to control their physicochemical properties is essential.14 The size, morphology, and surface charge of


3

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 4 of 34

nanoparticles, including liposomes, are well known to be important physicochemical parameters, which affect the therapeutic efficacy of pharmaceuticals based on these particles via their effect on the formulation properties.15–17 More recently, it has been recognized that the mechanical properties of nanoparticles also play an important role during in vitro and in vivo behaviors such as cellular uptake, blood circulation, and biodistribution.17 The popular experimental techniques for measuring the mechanical properties of liposomes are analysis of thermal fluctuation of the membrane, and measurement of the force needed to bend the membrane with micropipettes. However, these techniques are not applicable to nanosized liposomes because the techniques have to be performed under optical microscopy observation usually using cell-sized liposomes (at least 10 µm in diameter).18 Atomic force microscopy (AFM) is a direct method to not only observe the surface topology of nanoparticles immobilized on a solid substrate, but also to measure the mechanical property of their stiffness, by scanning the sample surface with an AFM cantilever.19 In this study, we refer to the slope of a force–deformation curve of a liposome as “stiffness”, which is a mechanical parameter interchangeable with bending modulus and Young’s modulus based on the theory of elasticity.20 By using a glass substrate coated with bovine serum albumin (BSA), we have developed a platform for the AFM measurement of nanosized liposomes in aqueous medium, and subsequently showed that their stiffness is a parameter affecting liposomal properties such as the release rate of encapsulated molecules, cellular uptake efficiency, and penetration efficiency into multicellular tumor spheroids.21-24 By AFM measurement, the stiffness of liposomes has been demonstrated to strongly depend on their thermodynamic phase-states, i.e., liquid disordered, liquid ordered, and solid ordered phases.22 To the best of our knowledge, there have been no quantitative studies of the effect of


4

Page 5 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

electrostatic interactions between the charged head groups of lipids on the stiffness of nanosized liposomes. However, such information may be crucial, since a liposomal bilayer membrane is an assembly of lipids balanced between, on one hand, the van der Waals and hydrophobic attractions of the acyl chains, and on the other hand, the electrostatic repulsion of the hydrophilic head groups.25 In this study, by means of AFM, we quantified the stiffness of nanosized liposomes composed of neutrally charged lipids, 1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) or 1,2-distearoylsn-glycero-3-phosphocholine (DSPC), combined with positively or negatively charged lipids while simultaneously imaging the liposomes in aqueous medium. To this end, our experimental technique using a BSA-coated glass substrate was applied to liposomes containing 1,2distearoyl-3-trimethylammoniumpropane

(DSTAP),

DOTAP,

1,2-dioleoyl-sn-glycero-3-

phospho-(1′-rac-glycerol) (DOPG), or DSPG. In addition, to discuss the usefulness of AFM, the stiffness data obtained by AFM were compared with Laurdan generalized polarization (GP) data, which are used to evaluate the hydration degree of nanosized liposomal membranes and often serve as an indicator of liposomal membrane fluidity.26

EXPERIMENTAL SECTION Materials. DOPC and BSA were purchased from Sigma-Aldrich (St. Louis, MO, USA). DSPC, DSTAP, DOTAP, and DOPG were purchased from Avanti Polar Lipids, Inc. (Alabaster, AL, USA). DSPG was purchased from Cayman Chemical Company (Ann Arbor, MI, USA). N,N-Dimethyl-6-dodecanoyl-2-naphthylamine (Laurdan) was purchased from Funakoshi Co., Ltd. (Tokyo, Japan). A slide glass (S-2411, 7S6 × 26 mm, thickness: 0.9–1.2 mm) and cover


5

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 6 of 34

glass (24 × 32 mm NEO micro cover glass) for AFM measurement were purchased from Matsunami Glass Ind., Ltd. (Osaka, Japan). All reagents were of analytical grade. Liposome preparation. Liposomes were prepared by a modification of a previously reported extrusion method.22 The lipid compositions of the liposomes used in this study are listed in Table 1, where the values in brackets of the lipid compositions represent %mol. Briefly, the total lipid (10 µmol) was mixed in chloroform in a round-bottomed flask, and the mixture was dried by evaporation to create a thin homogeneous lipid film. For the mixtures containing unsaturated lipids, the drying by evaporation was performed at room temperature. In contrast, for the mixtures containing saturated lipids, i.e., DSPC, DSTAP, or DSPG, the evaporation was performed at 70 °C. The lipid film was further dried overnight by means of vacuum desiccation to remove any residual solvent. The resultant lipid film was hydrated with 1 mL of 5% w/w glucose aqueous solution (pH 5.3) under mechanical agitation for 5 min either at 50 °C (for lipid films containing unsaturated lipids) or at 70 °C (for lipid films containing saturated lipids). The resultant dispersions were freeze-thawed 5 times using dry ice–methanol slush (−78 °C) and a water bath (50 °C for unsaturated lipids, 70 °C for saturated lipids), followed by extrusion 21 times (at room temperature for unsaturated lipids, 70 °C for saturated lipids) through a miniextruder (Avanti Polar Lipids) equipped with a 100-nm polycarbonate filter. All respective liposomes were obtained from three independent preparations. The polydispersity index values of the prepared liposomes, obtained by dynamic light scattering (DLS), were about 0.1, indicating monodispersity. Measurement of hydrodynamic diameter and zeta potential. DLS and laser Doppler velocimetry were performed at 25 °C using a Zetasizer Nano-ZS instrument equipped with the Zetasizer Software v.6.01 (Malvern Instruments, Malvern, UK). For the measurements, the


6

Page 7 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

prepared liposomal suspension was diluted with 5% w/w glucose aqueous solution (pH 5.3) so that the final concentration of total lipid was 0.2 mM. The hydrodynamic diameters and polydispersities of the liposomes were calculated by applying the cumulant method to the DLS data in the Zetasizer Software. The liposomal zeta potential was calculated from the velocimetry data by Henry’s approximation, according to the manufacturer’s guidance, in the software. Each measurement was performed in triplicate, and the measured values were averaged. The statistical measured values, i.e., means ± S.D., were obtained from three independent experimental results. Sample preparation for AFM experiment. For the AFM measurement of the liposomes in aqueous medium, we prepared the samples as follows. Using a biocompatible glue from JPK, a cover glass was glued onto a slide glass, and a plastic ring from JPK was glued to the cover glass surface. The container was rinsed with Milli-Q water and dried in air. The glass surface was coated with BSA according to our previous procedure.21,27 Then, 1.5 mL of 0.1–0.2 mM liposome suspension in 5% w/w glucose aqueous solution (pH 5.3) was incubated on the glass substrate for 20 min, followed by AFM measurement. We used 5% glucose solution for dispersion of liposomes owing to the instability of DOPC/DOTAP (50/50) liposomes in a physiological salt solution (phosphate buffered saline). We also incubated 1.5 mL of 0.1–0.2 mM DOPC/DOPG (50/50) liposome suspension in 5% w/w aqueous glucose solution (pH 7.4) on a BSA-coated glass substrate for 20 min. The liposome suspension was pre-equilibrated in the solution for 1 h prior to incubation. The pH of the 5% w/w glucose aqueous solution was adjusted to pH 7.4 by adding NaOH solution while the pH was monitored using a pH meter F-54. The final concentration of NaOH was ~0.02mM (HORIBA, Ltd., Kyoto, Japan). AFM measurement. To determine the stiffness of the liposomes, AFM measurement was performed at 25 ± 1 °C in aqueous medium by means of the QI mode™ of the JPK Nanowizard


7

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 8 of 34

Ultra Speed microscope equipped with JPK Data Processing Software v.6.0 (JPK Instruments AG, Berlin, Germany). The AFM piezo-scanner was calibrated with the manufacturer’s support. Prior to measurements, a soft cantilever (BioLever mini, nominal spring constant of 0.09 N/m, nominal tip radius < 8 nm, Olympus Co., Tokyo, Japan) was calibrated by the thermal noise method28,29 and was immersed in the sample for about 20 min to obtain thermal equilibrium. AFM images at a resolution of less than 8 nm/pixel were recorded with the following scanning parameters: set point, 0.1–15 nN; z-length, 80–100 nm; extend/retract speed, 15 µm/s. To correct for sample tilt, the obtained AFM images were subjected to flattening by the JPK Data Processing software. Images of the liposomes were then extracted using the Gwyddion software v.2.47, from which the maximum height and area-equivalent diameter of each liposome was determined.30 In the QI mode™ of the JPK Nanowizard Ultra Speed microscope, a force versus deformation curve was saved for each pixel of each AFM image,31 and the stiffness at the center of the liposome was analyzed by performing a linear fit in the linear region of each curve using the JPK Data Processing Software; the liposome stiffness corresponds to the value of the slope of the curves.20 Prior to collection of liposome stiffness data, we validated the cleanliness of our tips by monitoring the force curve at the substrate as described.32 Only liposomes with heights greater than about 35 nm were analyzed, since the measurement of objects with smaller heights is susceptible to interference by the solid substrate, as discussed in a review of AFM measurement of cells.33 For the measurement of each liposomal sample, the stiffness data were obtained for at least 10 liposomes, and averaged. The statistical values, i.e., means ± S.D., were obtained from three independent experiments.


8

Page 9 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

Evaluation of the hydration degree of liposomal membrane interface with Laurdan. The generalized polarization (GP) value of the fluorescent dye Laurdan embedded in the liposomal membranes was calculated from the fluorescence intensities by using the equation GP = (I440 − I490)/(I440 + I490), where I440 and I490 are the fluorescence intensities at emission wavelengths of 440 and 490 nm, respectively, under excitation at 340 nm.34-36 The fluorescence spectra of the samples were measured with an F-7000 fluorescence spectrophotometer (Hitachi HighTechnologies) at 25 ± 1°C. Samples were prepared by adding a stock solution of Laurdan in N,Ndimethylformamide to a liposome suspension to yield a lipid/Laurdan molar ratio of 100:1. The mixture was then incubated for 1 h in the dark at room temperature, and diluted in 5% w/w glucose aqueous solution (pH 5.3) so that the final concentrations of lipid and Laurdan in the test solution were 100 and 1 µM, respectively. To prepare liposome samples in 5% w/w glucose aqueous solution at pH 6.4, 7.4, and 8.0, the pH was measured with a pH cond meter F-54 (HORIBA, Ltd., Kyoto, Japan), and adjusted to the desired value by adding NaOH. Each measurement was performed in triplicate, and the measured values were averaged. The statistical measured values, i.e., means ± S.D., were obtained from three independent experimental results. Statistical analysis. Data were expressed as mean ± S.D. Results were analyzed by one-way analysis of variance with Bonferroni’s multiple comparison post-test. Differences were considered statistically significant at P < 0.05.

RESULTS AND DISCUSSION We investigated the liposome stiffness of DOPC (100), DOPC/DOTAP (50/50), and DOPC/DOPG (50/50) liposomes using a BSA-coated glass substrate in aqueous medium by


9

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 10 of 34

AFM (Figure 1). The zeta potential measurement showed that the surfaces of those three liposomes, respectively, are neutrally, positively, and negatively charged (Table 1). The calculation of bending modulus is based on finite element methods, or elastic models such as the Hertz, Sunnedon, or Shell models; however, the application of these models is inconsistent between researchers, and requires liposome information such as the elasticity in their deformation and the membrane thickness.20,37-40 Therefore, to exclude the potential errors resulting from using models, this study directly compared the stiffness data. It is known that liposomes have greater stiffness when they are smaller in size (i.e., when the liposomal membrane has higher curvature).20 The DLS data showed that the size distributions of DSPC/DSTAP (90/10) and DSPC/DSTAP (50/50) were slightly larger than those of other liposomes (Supporting Fig. S1A), being consistent with the result of Z-diameter of liposomes shown in Table 1. However, we confirmed that the liposomes analyzed by AFM all had similar height distribution (Supporting Fig. S1B). In addition, the average heights and area-equivalent diameters of liposomes analyzed by AFM showed no significant differences between the liposome samples (Supporting Table S1), thereby ruling out any variation in stiffness as a function of size. As shown in Figure 1A, the spherical structures of all three liposomes were observed to be adhered on the substrate; thus our AFM platform was applicable for the analysis of liposomes regardless of their surface charge. This successful charge-independent adhesion of liposomes onto a BSA-coated glass substrate implies that the adhesion is driven by non-electrostatic interactions as discussed in our previous studies.21,27 We then analyzed the force versus deformation curves obtained by AFM (Figure 1B and Figure 1C), and found that for the liposomes containing DOTAP or DOPG, the slopes of the curves were small (Figure 1C). As a


10

Page 11 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

result, the calculated stiffness values of DOPC/DOTAP (50/50) and DOPC/DOPG (50/50) liposomes are, respectively, 33% and 42% lower than that of DOPC (100) liposomes (Figure 1C). To investigate whether the charged head group of DOTAP and DOPG was responsible for the decrease in liposome stiffness, we further examined liposomes composed of saturated lipids (Figure 2). In this experiment, the DSPC (100) liposomes were replaced by DSPC/DSTAP (90/10), since DSPC (100) liposomes start aggregating immediately after preparation.41 Similarly to the case of liposomes composed of unsaturated lipids, the AFM images of the liposomes composed of DSPC/DSTAP (90/10), DSPC/DSTAP (50/50), or DSPC/DSPG (50/50) were successfully captured (Figure 2A). Likewise, the slopes of the force versus deformation curves of DSPC/DSTAP (50/50) and DSPC/DSPG (50/50) liposomes were found to be lower than that of DSPC/DSTAP (90/10) liposomes (Figures 2B and 2C). As a result, the calculated stiffness values of DSPC/DSTAP (50/50) and DSPC/DSPG (50/50) liposomes are, respectively, 41% and 59% lower than that of DSPC/DSTAP (90/10) liposomes (Figure 2C). To the best of our knowledge, this is the first study to demonstrate that charged lipids reduce the stiffness of nanosized liposomes independent of the saturation degree, and irrespective of whether the charge of the lipids is positive or negative. Strong electrostatic adhesion of a liposome onto a solid substrate decreases the membrane tension,42 which could lower the stiffness of the liposome; however, our result cannot be explained in terms of membrane tension alone. Indeed, although negatively charged DOPC/DOPG (50/50) and DSPC/DSPG (50/50) liposomes adhered to negatively charged BSAcoated glass substrates used in this study, where there is repulsive electrostatic interaction, the stiffness values are lower than those of their counterpart liposomes (Figures 1C and 2C).


11

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 12 of 34

Our finding implies that the decrease in stiffness of nanosized liposomes induced by charged lipids is derived from decreased bending energy via structural destabilization of the lipid bilayer membrane, which is induced by electrostatic repulsion between the charged head groups of lipids. Interestingly, we also found that the stiffness values of DOPC/DOPG (50/50) and DSPC/DSPG (50/50) liposomes were lower than those of DOPC/DOTAP (50/50) and DSPC/DSTAP (50/50) liposomes, respectively (Figures 1C and 2C). This may be because DOPG or DSPG caused greater destabilization of the liposomal membrane than that caused by DOTAP or DSTAP, because of the larger charge density of the phosphate group in the former compared with that of the trimethylammonium group in the latter. Previous studies using cell-sized liposomes or stacked multilamellar structures of lipids on a sample

chamber

have

shown

that

the

rigidity

of

lipid

membranes

containing

dioleoylphosphatidylserine (DOPS) is higher than that of those containing neutral lipids.43,44 These findings differ from our results. It should be noted that in the bilayer membrane system, the charged DOPS groups were highly protonated, at least below pH 5, or bound to counterions, resulting in “hard” domains with denser lipid packing via formation of hydrogen bonding between the lipid headgroups.43,44 A similar effect of ion binding can be observed in lipid headgroups of dipalmitoylphosphatidylglycerol.45 Even in supported lipid bilayers composed of neutral phosphatidylcholines on solid substrates, such membrane rigidification caused by ion binding has been confirmed at physiological salt concentration.46,47 Because we used 5% glucose solution in our system, owing to the instability of DOPC/DOTAP (50/50) liposomes in a physiological salt solution (phosphate buffered saline), this effect of ion binding is expected to have been small.


12

Page 13 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

The membrane curvature of the lipid bilayer must also be considered because the curvature of cell-sized liposomes is much lower than that of nanosized liposomes, which results in different packing densities and dynamics of the lipid molecules.48 A previous study reported that at low ion concentration, the bending rigidity of nanosized dimrystoilphosphatidylcholine (DMPC)liposomes significantly decreases when DOTAP or DOPC is incorporated, and the decrements are similar.49 Although the effect of DOTAP on liposomal membrane rigidity remains unknown, the length of the acylchain of DMPC is different to those of DOTAP and DOPC, which would lead to a lipid packing mismatch and a large decrease in the bending rigidity.18,49 To provide further insights into the effect of charged lipids on liposome stiffness, the stiffness was investigated in terms of the hydration degree of the liposomal membrane interface, as evaluated by the Laurdan GP method (Figures 3 and 4). Laurdan is known to homogeneously distribute in lipid membranes regardless of different coexisting lipid phases, and to locate at the hydrophilic–hydrophobic interface of the lipid bilayer, with the lauric acid tail anchored in the region of phospholipid acyl chains.35,36,50,51 Fluorescence spectra of Laurdan exhibit large Stokes shifts (the wavelength of maximum fluorescence typically shifts from ~440 nm to ~490 nm) resulting from water-induced dipolar relaxation of Laurdan from the excited state.35,36,51 Thus, the GP value contains information about the hydration degree of lipid interfaces, and is often used as an indicator of membrane stiffness because the hydration degree usually increases when the membrane fluidity is high, i.e., the membrane is soft.51-53 The fluorescence spectra of Laurdan incorporated into DOPC (100), DOPC/DOTAP (50/50), and DOPC/DOPG (50/50) liposome membranes are shown in Figure 3A. Regarding the GP values calculated from the fluorescence intensity at 440 nm relative to that at 490 nm, DOPC/DOTAP (50/50) and DOPC/DOPG (50/50) showed lower and higher GP, respectively, when compared with DOPC


13

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 14 of 34

(100). As a result, there was no correlation between GP and liposome stiffness (Figure 3B). The lower GP value of DOPC/DOTAP (50/50) is to be expected, since the charged lipid heads would contribute to electrostatic repulsion between lipid molecules and subsequently decrease the packing density of lipids in the liposomal membrane, thereby increasing the membrane hydration degree. Conversely, considering that this experiment was conducted in 5% w/w glucose aqueous solution at (acidic) pH 5.3, the larger GP value in DOPC/DOPG (50/50) than DOPC (100) may have been caused by the protonation of the phosphate group in DOPG (~1 < pKa < ~5). Indeed, previous studies have indicated the binding of positive counterions, such as sodium ions and protons, to the phosphate group in PG in liposomes containing egg yolk-derived PG, DOPG, or dipalmitoyl-PG.54-58 Therefore, the higher GP value of DOPC/DOPG (50/50) than DOPC (100) suggests that the PG-protonation decreases the electrostatic repulsion between lipid molecules in DOPC/DOPG (50/50) and results in a low degree of hydration, compared with DOPC (100) which has polar choline groups. To support this conclusion, the fluorescence spectra of Laurdan incorporated in DOPC/DOPG (50/50) were observed to be pH-dependent, namely, the GP value decreased with increasing pH (corresponding to reduced protonation) above pH 5 (Figures S1A and S1C). Despite the large difference in GP value between DOTAP and DOPG, the lack of difference in liposome stiffness observed between DOPC/DOTAP (50/50) and DOPC/DOPG (50/50) liposomes shown in Figure 3B suggests that the membrane rigidity of DOPC/DOPG (50/50) liposomes was reduced through repulsion of the charged head groups of the lipids, in addition to other factors. It is reported that a domain structure is formed by DMPG molecules,45 and that at the phase boundary regions of such nano-ordered domains, membrane thinning or lipid packing mismatch, or both, could disrupt membrane continuity and weaken the entire bilayer structure.18


14

Page 15 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

The presence of protonated PG groups in the liposomes might be a cause of the domain formation, leading to the weakness of the entire bilayer structure.45 To further investigate the lack of dependence of liposome stiffness on hydration, the stiffness of DOPC/DOPG (50/50) liposomes was also measured by AFM in 5% w/w aqueous glucose solution at pH 7.4 (Supporting Fig. S3), where the Laurdan GP value drops to −0.15 (Supporting Fig. S2C). As a result, the stiffness at pH 7.4 exhibited a similar value compared with the counterpart value at pH 5.3 (Supporting Fig. S3). This also shows that the hydration of the liposomal interface does not affect the stiffness, and stiffness is determined by other factors such as electrostatic repulsion between lipid molecules and the domain structures as described. This result supports the utility of AFM in assessing liposome stiffness when designing the lipid compositions of liposomes. No correlation between the Laurndan GP and the membrane fluidity in negatively charged liposomes composed of PC, phosphatidylethanolamine, and cardiolipin has been reported previously, which is consistent with our result.59 We further investigated the relationship between Laurdan GP and the stiffness of liposomes composed of saturated lipids, i.e., DSPC/DSTAP (90/10), DSPC/DSTAP (50/50), or DSPC/DSPG(50/50). Almost no difference could be observed between the fluorescence spectra of the three liposome samples (Figure 4A), and no correlation between the GP and the stiffness values was found (Figure 4B). In addition, no significant change in the Laurdan fluorescence spectra or the GP values of DSPC/DSPG (50/50) with varying pH was revealed, in contrast to the case of DOPC/DOPG (50/50) (Figures S1B and S1C). In general, for liposomes composed of saturated lipids, the Laurdan GP method has a poor sensitivity to changes in the hydration degree at the lipid interface, because of the much smaller change in the lipid packing state of saturated liposomes than that of liposomes composed of unsaturated lipids.34,60 AFM is therefore a


15

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 16 of 34

valuable alternative method to assess the stiffness of liposomes composed of saturated lipids where the hydration degree of the membrane cannot be evaluated by Laurdan.

CONCLUSIONS The present AFM study directly showed that charged lipids, whether negatively or positively charged, have the effect of reducing the stiffness of nanosized liposomes, independent of the saturation degree of the lipid acyl chains; the measured stiffness values of liposomes containing charged lipids are 30–60% lower than those of their neutral counterpart liposomes. The decrease in the liposome stiffness is considered to derive from mainly structural destabilization of the lipid bilayer membrane induced by electrostatic repulsion between the charged head groups of lipids. In addition to electrostatic repulsion between lipid molecules, other factors such as formation of a domain structure, which is formed depending on the lipid characteristics, might also affect the structural stability of lipid bilayers and liposome stiffness. In addition, we showed that the Laurdan GP values do not correlate with the AFM stiffness values of the liposomes prepared in this study, indicating that direct quantitative AFM is a more suitable method to gain molecularscale information about how the hydration degree of the liposomal interface reflects or does not reflect the liposome stiffness as a macroscopic property. Our AFM method will contribute to the quantitative characterization of nano-bio interaction, the optimization of the lipid composition of liposomes, and control of stiffness of liposomes containing charged lipids in order to develop liposomal formulations for clinical use.


16

Page 17 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

ASSOCIATED CONTENT Supporting Information The Supporting Information is available free of charge on the ACS Publications website at XXX. Heights and area-equivalent diameters of liposomes analyzed by AFM (Table S1). Size distribution of liposomes (Figure S1). Laurdan fluorescence measurement of liposomes in 5% w/w glucose aqueous solution at various pH (Figure S2). Comparison of stiffness of DOPC/DOPG (50/50) liposomes in 5% w/w aqueous glucose solution at pH 5.3 and 7.4 (Figure S3).

AUTHOR INFORMATION Corresponding Author *Email: [email protected] Notes The authors declare no competing financial interests.

ACKNOWLEDGMENTS This work was supported in part by the Research on Regulatory Harmonization and Evaluation of Pharmaceuticals, Medical Devices, Regenerative and Cellular Therapy Products, Gene Therapy Products, and Cosmetics from the Japan Agency for Medical Research and Development, AMED (17mk0101082j0101).


17

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 18 of 34

REFERENCES (1)

Sainz, V.; Conniot, J.; Matos, A. I.; Peres, C.; Zupannčič, E.; Moura, L.; Silva, L. C.;

Florindo, H. F.; Gaspar, R. S. Regulatory Aspects on Nanomedicines. Biochem. Biophys. Res. Commun. 2015, 468, 504–510. (2)

Bozzuto, G.; Molinari, A. Liposomes as Nanomedical Devices. Int. J. Nanomedicine

2015, 10, 975–999. (3)

Adler-moore, J. P.; Proffitt, R. T. Development, Characterization, Efficacy and Mode of

Action of Ambisome, A Unilamellar Liposomal Formulation of Amphotericin B. J. Liposome Res. 1993, 3, 429–450. (4)

Oh, Y. K.; Nix, D. E.; Straubinger, R. M. Formulation and Efficacy of Liposome-

Encapsulated Antibiotics for Therapy of Intracellular Mycobacterium Avium Infection. Antimicrob. Agents Chemother. 1995, 39, 2104–2111. (5)

Amin, K.; Ng, K. Y.; Brown, C. S.; Bruno, M. S.; Heath, T. D. LDL Induced Association

of Anionic Liposomes with Cells and Delivery of Contents as Shown by the Increase in Potency of Liposome Dependent Drugs. Pharm. Res. 2001, 18, 914–921. (6)

Nie, Y.; Ji, L.; Ding, H.; Xie, L.; Li, L.; He, B.; Wu, Y.; Gu, Z. Cholesterol Derivatives

Based Charged Liposomes for Doxorubicin Delivery: Preparation, in Vitro and in Vivo Characterization. Theranostics 2012, 2, 1092–1103. (7)

Gjetting, T.; Arildsen, N. S.; Christensen, C. L.; Poulsen, T. T.; Roth, J. A.; Handlos, V.

N.; Poulsen, H. S. In Vitro and in Vivo Effects of Polyethylene Glycol (PEG)-modified Lipid in DOTAP/Cholesterol-mediated Gene Transfection. Int. J. Nanomedicine 2010, 5, 371–383.


18

Page 19 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

(8)

Nogueira, E.; Gomes, A. C.; Preto, A.; Cavaco-Paulo, A. Design of Liposomal

Formulations for Cell Targeting. Colloids Surfaces B: Biointerfaces 2015, 136, 514–526. (9)

Majzoub, R. N.; Wonder, E.; Ewert, K. K.; Kotamraju, V. R.; Teesalu, T.; Safinya, C. R.

Rab11 and Lysotracker Markers Reveal Correlation between Endosomal Pathways and Transfection Efficiency of Surface-Functionalized Cationic Liposome-DNA Nanoparticles. J. Phys. Chem. B 2016, 120, 6439–6453. (10) Lee, J.; Kim, J.; Jeong, M.; Lee, H.; Goh, U.; Kim, H.; Kim, B.; Park, J. H. LiposomeBased Engineering of Cells to Package Hydrophobic Compounds in Membrane Vesicles for Tumor Penetration. Nano Lett. 2015, 15, 2938–2944. (11) Dakwar, G. R.; Shariati, M.; Willaert, W.; Ceelen, W.; De Smedt, S. C.; Remaut, K. Nanomedicine-Based Intraperitoneal Therapy for the Treatment of Peritoneal CarcinomatosisMission Possible? Adv. Drug Deliv. Rev. 2017, 108, 13–24. (12) Raviv, U.; Needleman, D. J.; Li, Y.; Miller, H. P.; Wilson, L.; Safinya, C. R. Cationic Liposome-Microtubule Complexes: Pathways to the Formation of Two-State Lipid-Protein Nanotubes with Open or Closed Ends. Proc. Natl. Acad. Sci. U. S. A. 2005, 102, 11167–11172. (13) Regelin, A. E.; Fankhaenel, S.; Gürtesch, L.; Prinz, C.; Von Kiedrowski, G.; Massing, U. Biophysical and Lipofection Studies of DOTAP Analogs. Biochim. Biophys. Acta 2000, 1464, 151–164. (14) Barenholz, Y. Doxil Ⓡ -The First FDA-approved Nano-Drug: Lessons Learned. J. Control. Release 2012, 160, 117–134.


19

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 20 of 34

(15) Ehmann, F.; Sakai-Kato, K.; Duncan, R.; Perez, de la Ossa D H.; Pita, R.; Vidal, J-M.; Kohli A.; Tothfalusi, L; Sanh, A.; Tinton, S.; Robert, J.-L. ; Lima, B.S.; Amati, M.P. Nextgeneration Nanomedicines and Nanosimilars: EU Regulators' Initiatives Relating to the Development and Evaluation of Nanomedicines. Nanomedicine 2013, 8, 849-856. (16) Sakai-Kato, K.; Nishiyama, N.; Kozaki, M.; Nakanishi, T.; Matsuda, Y.; Hirano, M.; Hanada, H.; Hisada, S.; Onodera, H.; Harashima, H.; Matsumura, Y.; Kataoka, K.; Goda, Y.; Okuda, H.; and Kawanishi, T. General Considerations Regarding the in Vitro and in Vivo Properties of Block Copolymer Micelle Products and their Evaluation. J. Control. Release 2015, 210, 76-83. (17) Anselmo, A. C.; Mitragotri, S. Impact of Particle Elasticity on Particle-based Drug Delivery Systems. Adv. Drug Deliv. Rev. 2017, 108, 51–67. (18) Dimova, R. Recent Developments in the Field of Bending Rigidity Measurements on Membranes. Adv. Colloid Interface Sci. 2014, 208, 225–234. (19) Maver, U.; Velnar, T.; Gaberšček, M.; Planinšek, O.; Finšgar, M. Recent Progressive Use of Atomic Force Microscopy in Biomedical Applications. Trends Anal. Chem. 2016, 80, 96–111. (20) Delorme, N.; Fery, A. Direct Method to Study Membrane Rigidity of Small Vesicles Based on Atomic Force Microscope Force Spectroscopy. Phys. Rev. E 2006, 74, 30901. (21) Takechi-Haraya, Y.; Sakai-Kato, K.; Abe, Y.; Kawanishi, T.; Okuda, H.; Goda, Y. Observation of Liposomes of Differing Lipid Composition in Aqueous Medium by Means of Atomic Force Microscopy. Microscopy 2016, 65, 383–389.


20

Page 21 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

(22) Takechi-Haraya, Y.; Sakai-Kato, K.; Abe, Y.; Kawanishi, T.; Okuda, H.; Goda, Y. Atomic Force Microscopic Analysis of the Effect of Lipid Composition on Liposome Membrane Rigidity. Langmuir 2016, 32, 6074–6082. (23) Takechi-Haraya, Y.; Sakai-Kato, K.; Goda, Y. Membrane Rigidity Determined by Atomic Force Microscopy Is a Parameter of the Permeability of Liposomal Membranes to the Hydrophilic Compound Calcein. AAPS PharmSciTech 2017, 18, 1887–1893. (24) Takechi-Haraya, Y.; Goda, Y.; Sakai-Kato, K. Control of Liposomal Penetration into Three-Dimensional Multicellular Tumor Spheroids by Modulating Liposomal Membrane Rigidity. Mol. Pharm. 2017, 14, 2158–2165. (25) May, S. A Molecular Model for the Line Tension of Lipid Membranes. Eur. Phys. J. E 2000, 44, 37–44. (26) Soto-Arriaza, M. A; Sotomayor, C. P.; Lissi, E. A. Relationship between Lipid Peroxidation and Rigidity in L-Alpha-Phosphatidylcholine-DPPC Vesicles. J. Colloid Interface Sci. 2008, 323, 70–74. (27) Takechi-Haraya, Y.; Goda, Y.; Sakai-Kato, K. Imaging and Size Measurement of Nanoparticles in Aqueous Medium by Use of Atomic Force Microscopy. Anal. Bioanal. Chem. 2018, 410, 1525–1531. (28) Hutter, J. L.; Bechhoefer, J. Calibration of Atomic-Force Microscope Tips. Rev. Sci. Instrum. 1993, 64, 1868–1873. (29) Sader, J. E.; Chon, J. W. M.; Mulvaney, P. Calibration of Rectangular Atomic Force Microscope Cantilevers. Rev. Sci. Instrum. 1999, 70, 3967–3969.


21

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 22 of 34

(30) Nečas, D.; Klapetek, P. Gwyddion: An Open-Source Software for SPM Data Analysis. Cent. Eur. J. Phys. 2012, 10, 181–188. (31) Smolyakov, G.; Formosa-Dague, C.; Severac, C.; Duval, R. E.; Dague, E. High Speed Indentation Measures by FV, QI and QNM Introduce a New Understanding of Bionanomechanical Experiments. Micron 2016, 85, 8–14. (32) Vorselen, D.; Mackintosh, F. C.; Roos, W. H.; Wuite, G. J. L. Competition between Bending and Internal Pressure Governs the Mechanics of Fluid Nanovesicles. ACS Nano 2017, 11, 2628–2636. (33) Kuznetsova, T. G.; Starodubtseva, M. N.; Yegorenkov, N. I.; Chizhik, S. A.; Zhdanov, R. I. Atomic Force Microscopy Probing of Cell Elasticity. Micron 2007, 38, 824–833. (34) Kitayama, H.; Takechi, Y.; Tamai, N.; Matsuki, H.; Yomota, C.; Saito, H. Thermotropic Phase Behavior of Hydrogenated Soybean Phosphatidylcholine-Cholesterol Binary Liposome Membrane. Chem. Pharm. Bull. 2014, 62, 58–63. (35) Parasassi, T.; Krasnowska, E. K.; Bagatolli, L.; Gratton, E. Laurdan and Prodan as Polarity-Sensitive Fluorescent Membrane Probes. J. Fluoresc. 1998, 8, 365–373. (36) De Vequi-Suplicy, C. C.; Benatti, C. R.; Lamy, M. T. Laurdan in Fluid Bilayers: Position and Structural Sensitivity. J. Fluoresc. 2006, 16, 431–439. (37) Liang, X.; Mao, G.; Ng, K. Y. S. Mechanical Properties and Stability Measurement of Cholesterol-Containing Liposome on Mica by Atomic Force Microscopy. J. Colloid Interface Sci. 2004, 278, 53–62.


22

Page 23 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

(38) Ramachandran, S.; Quist, A. P.; Kumar, S.; Lal, R. Cisplatin Nanoliposomes for Cancer Therapy: AFM and Fluorescence Imaging of Cisplatin Encapsulation, Stability, Cellular Uptake, and Toxicity. Langmuir 2006, 22, 8156–8162. (39) Li, S.; Eghiaian, F.; Sieben, C.; Herrmann, A.; Schaap, I. A. T. Bending and Puncturing the Influenza Lipid Envelope. Biophys. J. 2011, 100, 637-645. (40)

Et-Thakafy, O.; Delorme, N.; Gaillard, C.; Mériadec, C.; Artzner, F.; Lopez, C.;

Guyomarch, F. Mechanical Properties of Membranes Composed of Gel-Phase or Fluid-Phase Phospholipids Probed on Liposomes by Atomic Force Spectroscopy. Langmuir 2017, 33, 5117– 5126. (41) Dos Santos, N.; Allen, C.; Doppen, A. M.; Anantha, M.; Cox, K. A. K.; Gallagher, R. C.; Karlsson, G.; Edwards, K.; Kenner, G.; Samuels, L.; Webb, M. S.; Bally, M. B. Influence of Poly(ethylene glycol) Grafting Density and Polymer Length on Liposomes: Relating Plasma Circulation Lifetimes to Protein Binding. Biochim. Biophys. Acta 2007, 1768, 1367–1377. (42) Murrell, M. P.; Voituriez, R.; Joanny, J. F.; Nassoy, P.; Sykes, C.; Gardel, M. L. Liposome Adhesion Generates Traction Stress. Nat. Phys. 2014, 10, 163–169. (43) Petrache, H. I.; Tristram-nagle, S.; Gawrisch, K.; Harries, D.; Parsegian, V. A.; Nagle, J. F. Structure and Fluctuations of Charged Phosphatidylserine Bilayers in the Absence of Salt. Biophys. J. 2004, 86, 1574–1586. (44) Mitkova, D.; Marukovich, N.; Ermakov, Y. A.; Vitkova, V. Bending Rigidity of Phophatidylserine-containing lipid bilayers in acidic aqueous solutions. Colloids and Surfaces A: Physicochem. Eng. Aspects 2014, 460, 71–78.


23

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 24 of 34

(45) Garidel, P.; Johann, C.; Mennicke, L.; Blume, A. The Mixing Behavior of Pseudobinary Phosphatidylcholine-phosphatidylglycerol Mixtures as a Function of pH and Chain Length. Eur. Biophys. J. 1997, 26, 447–459. (46) Garcia-Manyes, S.; Sanz, F. Nanomechanics of Lipid Bilayers by Force Spectroscopy with AFM: A Perspective. Biochim. Biophys. Acta 2010, 1798, 741–749. (47) Redondo-Morata, L.; Oncins, G.; Sanz, F. Force Spectroscopy Reveals the Effect of Different Ions in the Nanomechanical Behavior of Phospholipid Model Membranes: The Case of Potassium Cation. Biophys. J. 2012, 102, 66–74. (48)

Takechi, Y.; Saito, H.; Okamura, E. Slow Tumbling but Large Protrusion of

Phospholipids in the Cell Sized Giant Vesicle. Chem. Phys. Lett. 2013, 570, 136–140. (49) Büning, B.; Stehle, R.; Falus, P.; Farago, B. Influence of Charge Density on Bilayer Bending Rigidity in Lipid Vesicles : A Combined Dynamic Light Scattering and Neutron SpinEcho Study. Eur. Phys. J. E 2013, 36, 77. (50) Bagatolli, L. A. To See or Not to See: Lateral Organization of Biological Membranes and Fluorescence Microscopy. Biochim. Biophys. Acta 2006, 1758, 1541–1556. (51) Parasassi, T.; De Stasio, G.; Ravagnan, G.; Rusch, R. M.; Gratton, E. Quantitation of Lipid Phases in Phospholipid Vesicles by the Generalized Polarization of Laurdan Fluorescence. Biophys. J. 1991, 60, 179–189. (52) Parasassi, T.; Di Stefano, M.; Loiero, M.; Ravagnan, G.; Gratton, E. Cholesterol Modifies Water Concentration and Dynamics in Phospholipid Bilayers: A Fluorescence Study Using Laurdan Probe. Biophys. J. 1994, 66, 763–768.


24

Page 25 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

(53) Parasassi, T.; Gratton, E.; Yu, W. M.; Wilson, P.; Levi, M. Two-Photon Fluorescence Microscopy of Laurdan Generalized Polarization Domains in Model and Natural Membranes. Biophys. J. 1997, 72, 2413–2429. (54) Khalifat, N.; Rahimi, M.; Bitbol, A.; Seigneuret, M.; Fournier, J.; Puff, N.; Arroyo, M.; Angelova, M. I. Interplay of Packing and Flip-Flop in Local Bilayer Deformation . How Cardiolipin-Deficient Yeast Mutant. Biophysj 2014, 107, 879–890. (55) Tocanne, J.-F.; Teissié, J. Ionization of Phospholipids and Phospholipid-Supported Interfacial Lateral Diffusion of Protons in Membrane Model Systems. Biochim. Biophys. Acta 1990, 1031, 111–142. (56) Kubsch, B.; Robinson, T.; Lipowsky, R.; Dimova, R. Solution Asymmetry and Salt Expand Fluid-Fluid Coexistence Regions of Charged Membranes. Biophys. J. 2016, 110, 2581– 2584. (57) Garidel, P.; Blume, A. Miscibility of Phosphatidylethanolamine-Phosphatidylglycerol Mixtures as a Function of pH and Acyl Chain Length. Eur. Biophys. J. 2000, 28, 629–638. (58) Filippov, A.; Orädd, G.; Lindblom, G. Effect of NaCl and CaCl2 on the Lateral Diffusion of Zwitterionic and Anionic Lipids in Bilayers. Chem. Phys. Lipids 2009, 159, 81–87. (59) Khalifat, N.; Fournier, J.; Angelova, M. I.; Puff, N. Lipid Packing Variations Induced by pH in Cardiolipin-Containing Bilayers: The Driving Force for the Cristae-like Shape Instability. Biochim. Biophys. Acta 2011, 1808, 2724–2733.


25

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 26 of 34

(60) Matsuki, H.; Goto, H.; Kusube, M.; Tamai, N. Imaging of Phosphatidylcholine Bilayers by a High-Pressure Fluorescence Technique: Detection of the Packing Difference. Bull. Chem. Soc. Jpn. 2011, 84, 1329–1335.


26

Page 27 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

Table 1. Zeta potentials and hydrodynamic diameters of liposomes

Lipid composition

Zeta potential (mV)

Diameter (nm)

0.62 ± 5.0

113 ± 1.7

80 ± 8.3

109 ± 5.0

−107 ± 3.1

116 ± 1.7

DSPC/DSTAP (90/10), saturatedb

70 ± 7.6

130 ± 13

DSPC/DSTAP (50/50), saturatedb

85 ± 6.0

143 ± 7.5

DSPC/DSPG (50/50), saturatedb

-77 ± 5.3

116 ± 6.1

DOPC (100), unsaturateda DOPC/DOTAP (50/50), unsaturateda DOPC/DOPG (50/50), unsaturateda

a

Liposomes composed of unsaturated lipids. bLiposomes composed of saturated lipids. All values are means ± S.D (n = 3). The values in brackets are %mol. DOPC, 1,2-dioleoyl-snglycero-3-phosphocholine; DOTAP, 1,2-dioleoyl-3-trimethylammoniumpropane; DOPG, 1,2dioleoyl-sn-glycero-3-phospho-(1′-rac-glycerol); DSPC, 1,2-distearoyl-sn-glycero-3phosphocholine; DSTAP, 1,2-distearoyl-3-trimethylammoniumpropane; DSPG, 1,2-distearoylsn-glycero-3-phospho-(1′-rac-glycerol).


27

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 28 of 34

FIGURE CAPTIONS Figure 1. AFM measurement of DOPC (100), DOPC/DOTAP (50/50), and DOPC/DOPG (50/50) liposomes in 5% w/w glucose aqueous solution. (A) AFM images of liposomes on BSAcoated glass. The scale bar in the lower right of each AFM image represents 200 nm. Z-scales are shown in the right of each AFM image. (B) Force–deformation curves at the center of liposomes on BSA-coated glass. The inset shows the force–deformation curve as a reference obtained at the substrate. (C) Liposomal stiffness of liposomes. Total number (N) of analyzed liposomes: N = 152 for DOPC (100), N = 290 for DOPC/DOTAP (50/50), N = 52 for DOPC/DOPG (50/50) liposomes. **P < 0.01, compared with DOPC (100) liposomes. n.s., not significant. Figure 2. AFM measurement of DSPC/DSTAP (90/10), DSPC/DSTAP (50/50), and DSPC/DSPG (50/50) liposomes in 5% w/w glucose aqueous solution. (A) AFM images of liposomes on BSA-coated glass. The scale bar in the lower right of each AFM image represents 200 nm. Z-scales are shown in the right of each AFM image. (B) Force–deformation curves at the center of liposomes on BSA-coated glass. The inset shows the force–deformation curve as a reference obtained at the substrate. (C) Liposomal stiffness of liposomes. Total number (N) of analyzed liposomes: N = 105 for DSPC/DSTAP (90/10), N = 117 for DSPC/DSTAP (50/50), N = 205 for DSPC/DSPG (50/50) liposomes. **P < 0.01, compared with DSPC/DSTAP (90/10) liposomes. n.s., not significant. Figure 3. (A) Fluorescence spectra of Laurdan in DOPC (100), DOPC/DOTAP (50/50), and DOPC/DOPG (50/50) liposomes in 5% w/w glucose aqueous solution. (B) Plots of stiffness values against Laurdan GP values in liposomes.


28

Page 29 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir

Figure 4. (A) Fluorescence spectra of Laurdan in DSPC/DSTAP (90/10), DSPC/DSTAP (50/50), and DSPC/DSPG (50/50) liposomes in 5% w/w glucose aqueous solution. (B) Plots of stiffness values against Laurdan GP values in liposomes.


29

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Page 30 of 34

30

Page 31 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir


31

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Page 32 of 34

32

Page 33 of 34 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Langmuir


33

Langmuir 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 34 of 34

Table of Contents (area no larger than 8.5 cm × 4.75 cm)


34

Atomic force microscopy study on stiffness of nanosized liposomes

Recommend Documents