Low-Frequency Ultrasound-Induced Transport across Non-Raft

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Low Frequency Ultrasound-Induced Transport Across Non-Raft-Forming Ternary Lipid Bilayers Eleanor Frances Small, Nily R. Dan, and Steven P Wrenn Langmuir, Just Accepted Manuscript • Publication Date (Web): 13 Sep 2012 Downloaded from http://pubs.acs.org on September 26, 2012

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Low Frequency Ultrasound-Induced Transport Across Non-Raft-Forming Ternary Lipid Bilayers

Eleanor F. Small, Nily R. Dan, Steven P. Wrenn* Department of Chemical and Biological Engineering, Drexel University, 3141 Chestnut Street, Philadelphia, PA 19104

ABSTRACT

We examined the effect of bilayer composition on membrane sensitivity to low frequency ultrasound (LFUS) in bilayers composed of ternary mixtures of 1-palmitoyl-2-oleoyl-sn-glycero3-phosphocholine (POPC), dipalmitoyl-phosphocholine (DPPC), and cholesterol. The phase diagram of this system does not display macroscopic phase coexistence between liquid phases (although there are suggestions that there is coexistence between a liquid and a solid phase). Samples from across the composition space were exposed to 20 kHz, continuous wave ultrasound and the response of the bilayer was quantified using steady-state fluorescence spectroscopy to measure the release of a self-quenching dye, calcein, from large unilamellar vesicles. Dynamic light scattering measurements indicate that, in this system, release proceeds

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primarily by transport through the vesicle bilayer. While vesicle destruction might account – at least in part – for the light scattering trends observed, evidence of destruction was not as obvious as in other lipid systems. Values for bilayer permeability are obtained by fitting release kinetics to a two-film theory mathematical model. The permeability due to LFUS is found to increase with increasing DPPC content, as the bilayer tends toward the solid-ordered phase. Permeability, and thus sensitivity to LFUS, decreases with either POPC or cholesterol mole fractions. In the liquid regime of this system there is no recorded phase transition, thus cholesterol is the determining factor in release rates. However, the presence of domain boundaries between distinctly differing phases of liquid and solid is found to cause release rates to more than double. The correlation of permeability with phase behavior might prove useful in designing and developing therapies based on ultrasound and membrane interactions.

Introduction Synthetic lipid bilayers have been extensively studied, both as a model system representing aspects of cell membranes,1-5 and for applications ranging from cosmetics to drug delivery.6-8 These lipid vesicles, referred to hereafter as liposomes, have ideal encapsulation abilities (hydrophobic and hydrophilic load capabilities)9 and are easily functionalized for prolonged circulation in the body,8 but triggering drug release has been the primary challenge to their successful use.10,11 Low frequency ultrasound (LFUS) has been found to enhance the permeability of biological membranes,8,9,12,13 as well as induce release of encapsulated molecules from liposomes in vitro.10,14-16 Unlike high frequency ultrasound, where the release mechanism was linked to


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heating effects17-20 or microbubble cavitation,21-24 LFUS has been found to increase the permeability of lipid membranes through non-thermal effects.9,16,25 In the synthetic systems, the effects of LFUS were found to be transient; once sonication ceased, so did the release.14,16,25 Analysis of LFUS-induced release from liposomes suggests that it is dominated by a diffusive process, concurrent with some contribution from liposome destruction.26,27 Both processes can be linked to changes in lipid packing in the membrane: Indeed, Mendelsohn et al.28 found that US disrupts the packing of the hydrophobic lipid tails in the bilayer core. LFUS was shown to create structural defects in liposomes at a temperature below their phase transition, resulting in enhanced permeation.29 The leading theory for low frequency ultrasound-induced leakage is pore formation;8,16,25,30,31 however, the detailed mechanism by which such pores form is not known and remains an active area of investigation.11,21,32,33 The rate of LFUS-induced release has been linked to US characteristics (frequency, intensity),8 as well as liposome size,25 lipid type,10,34 and addition of surfactants to the membrane.14,25,30 These studies produced only qualitative observations concerning the relationship between membrane phase behavior and liposome leakage. Three component lipid mixtures, where one of the components is cholesterol, have been found to display different phases and a rich variety of complex phase coexistence that depend on the type of lipid constituents.35-39 Recently, we investigated the effect of bilayer phase behavior on sensitivity to LFUS in large unilamellar vesicles composed of 1,2-dioleoyl-phosphocholine (DOPC), 1,2-dipalmitoyl-phosphocholine (DPPC), and cholesterol.27 DPPC is a high melting temperature, saturated lipid, while DOPC has a relatively low melting point and unsaturated acyl chains. Several studies of the phase behavior of DOPC/DPPC/cholesterol bilayers find a region of macroscopic phase coexistence between the two liquid-phases (liquid-disordered, ld and liquid-ordered, lo) with well-defined tie lines.38,40


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Solid phases (so) were found to nucleate from the liquid-ordered phases in mixtures where the cholesterol mole fraction was lower than 0.1.37,38 The sensitivity of the bilayer to LFUS-induced leakage was determined by measuring the rate of release of self-quenching, encapsulated calcein;27 more sensitive bilayers would display faster release (either through bilayer permeation, vesicle destruction, or a combination of both). We found that the rate of release was not set by the cholesterol content (mole fraction) per se but by the lipid-phase. Bilayers with identical cholesterol mole fraction but a different phase state (due to different DOPC to DPPC ratios) displayed significant differences in the release rate, and thus in sensitivity to LFUS. Generally, the rate of release was found to be faster in liquid-disordered phases than in liquid-ordered phases, with intermediate values in the two-phase coexistence region.27 The DOPC mixture studies represent one type of lipid phase diagrams. Another class of lipid mixtures shows very different phase behavior with no macroscopic coexistence, or clear transition, between the ld and lo phases.39,41,42 These lipid mixes were also found to display macroscopic phase coexistence between the liquid-disordered and the gel, or solid, phase, ldso.39,43 Thus, unlike the DOPC system, where the macroscopic phase coexistence indicated a 1st order phase transition, these lipid mixtures may represent continuous phase transitions, which can greatly affect their functions.41,44 In this paper we investigate the effect of membrane phase on LFUS sensitivity in such a lipid system, comprising mixtures of 1-palmitoyl-2-oleoyl-snglycero-3-phosphocholine (POPC) with DPPC and cholesterol. Unlike the DOPC-based system, the POPC-based system under investigation here displays no proven liquid-liquid coexistence region


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Although the POPC system falls into the category of phase diagrams with no macroscopic phase coexistence between liquid phases,37,41 it does display regions with local concentration inhomogeneities.36,41 Brown et al.36 used polarity-dependent emission maximum shift of DANPC to detect the type of liquid phase(s) present in the membrane. The fluorescent probe partitions preferentially into the liquid-disordered phase and displays a shift in the emission wavelength when forced into the liquid-ordered phase. The shift in environment polarity causes a blue shift in DANSYL max emission wavelength.45 A sharp blue shift in DAN-PC was observed in the POPC/DPPC/Cholesterol system. The DAN-PC spectral shift results in Brown et al.36 indicate two different phases exist in the system even if there is no observable region of coexistence. Since the transition from ld to lo could not be determined from macroscopic phase-separation studies, we mapped the transition using the results of Brown et al.36 This paper seeks to build on previous work by quantifying the relationship between membrane composition and ultrasound-induced release from liposomes with continuous phase transition lipid mixtures. We present another “release map”, in which measured leakage results are superimposed onto a phase diagram. Additionally, kinetic release profiles are fit with simple two-film theory mathematical models describing diffusion and bilayer destruction. Quite surprisingly, we do not find a consistent correlation between the rate of release and the liquid phase type; samples in the expected liquid-ordered regime did not display release rates (and thus, susceptibility to LFUS) that were consistently lower than those of samples in the expected liquiddisordered state. Moreover, the most rapid release rates, and therefore response to LFUS, occurred in samples that are in the ld - so phase coexistence regime. These results suggest that for continuous phase transition lipid mixtures studied here, LFUS response is not sensitive to the


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type of liquid phase but rather to the presence of domain boundaries between distinctly differing phases.

Materials and Methods Materials. Samples of 1-palmitoyl-2-oleoyl-phosphatidylcholine (POPC), and 1, 2dipalmitoyl-phosphocholine (DPPC) were provided by Lipoid, LLC (Newark, NJ) or purchased from Avanti Lipids (Alabaster, AL). Cholesterol (Chol), sodium chloride (NaCl), Calcein, Triton X-100, Sephadex G-50, Sephadex G-75, and ethylenediaminetetra-acetic acid (EDTA anhydrous) were purchased from Sigma-Aldrich (St. Louis, MO).

TrisBase (Tris) and

chloroform were purchased from Fisher Scientific, Inc. (Fair Lawn, NJ). Extrusion drain disks and poly carbon filters were purchased from Nuclepore, Whatman Inc. (Clifton, NJ). Nitrogen (N2) was obtained from Airgas (Allentown, PA).

All products were used without further

purification. Vesicle Preparation. Samples for both “release map” and kinetic studies were created along lines of constant DPPC (5, 20, 40, 60, and 80%) with varying cholesterol compositions, in intervals of 5%, between 5-60%. Large unilamellar vesicles (LUVs), nominally 200nm in diameter, were created by extrusion of multilamellar vesicles (MLVs) to maintain consistency with research reviewed above and with previous work by this lab. Multilamellar vesicles were created using the Rapid Solvent Exchange (RSE) method,46,47 to avoid undesired cholesterol crystallization at the higher cholesterol samples. A detailed procedure for liposome creation was described in our previous work.27 Calcein Buffer (150mM NaCl, 10mM Tris, 1mM EDTA, and 70mM Calcein, pH 7.4) was encapsulated in the liposomes and samples were subject to size exclusion chromatography to separate the vesicles from un-encapsulated calcein. Elution buffer


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was room temperature Ultrasound Buffer (150mM NaCl, 10mM Tris, and 1mM EDTA, at pH 7.4). Column eluate was collected and divided in to three, 3 mL samples and analyzed using a Brookhaven 90 Plus, Dynamic Light Scattering (DLS) apparatus. For each composition one of the three samples was selected as a control, in which fluorescence was measured periodically but no sonication was applied. The other aliquots, referred to as “test samples”, were then exposed to ultrasound and rest cycles as described below. Analysis of samples on DLS obtained effective diameter and polydispersity statistics. At all lipid compositions the average vesicle diameter was 202 ± 18nm. DLS and Release Fluorescence Assays. All release assays were performed using a Misonix XL2020 probe tip sonicator (Misonix Inc., Farmingdale, NY) operating at 20 kHz using a 419 tip. The Sonicator was calibrated using the Misonix tuning protocol and set to 10% output power (electrical) at a setting of 3. The peak-to-peak pressure amplitudes and spatial-peak, temporal-peak intensity were measured using a Reson TC4038 hydrophone (Reson Inc., US, Goleta, CA) with a sensitivity of -228.2 dB re 1V/µPa at 20 kHz. To protect the hydrophone the acoustic pressure amplitude measurements were performed at 10, 9, and 8cm radial distances from the probe’s tip, measurement data were used to estimate the pressure amplitude at 1cm (the distance at which release assays were performed). At the given setting, the amplitude was calculated to be equal to approximately 256 ± 26 kPa, which corresponds to a spatial-peak, temporal-peak intensity of approximately 2.2W/cm2. Fluorescence studies used a well-established assay of relief of self-quenching of calcein. Release and destruction assays are described in detail in our previous work.27 Briefly “Release map” data were gathered by exposing the test sample to 180 consecutive seconds of continuous


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wave, 20 kHz ultrasound. Kinetic release studies were performed by exposing the test sample to 30 second intervals of continuous wave ultrasound, with a three minute rest period between ultrasonic exposures, until total ultrasonic exposure reached 360 seconds. A steady-state fluorescence spectrometer (Photon Technology International, Ontario, Canada, model Q-5/W601) measured fluorescence after each ultrasound exposure. Samples were placed in an ice water bath to maintain their temperature at 22 ± 2°C. In all cases samples reached at least 60% release. Fraction of release was determined via: ( It − I t =0 ) /( I max − I t =0 ) where It represents the intensity of the current run, It=0 is the intensity prior

to any ultrasonic exposure (t=0), and Imax is the intensity after adding 10µl of TritonX-100 (final concentration of 5.7E-3M), which destroys all the membranes releasing all encapsulated dye 14,25,48,49

. Triton X-100 had negligible effect on calcein fluorescence at the concentration used.27

Samples were excited at 488nm and peak emission was found to be 521 ± 3nm. Destruction studies were performed using vesicles prepared with Ultrasound Buffer. Samples were then subject to initial DLS measurements followed by kinetic study-style sonication. DLS readings were taken after every 120 seconds of LFUS exposure, including a final reading after 360 seconds. “Destruction” in this case, as in our previous work,27 is defined as catastrophic damage to the bilayer causing it to lose vesicle formation. Molecular dynamics simulations by Leontiadou et al.50 support theories suggesting the possibility of vesicle destruction. Their simulations revealed a critical tension above which LFUS-created transient pores become unstable and the membrane ruptures. Modeling. Model parameter 1/τ, for destruction-only limit, was fit using the built-in least squares fit function “Lsqcurvefit” in MatLAB R2010a (The MathWorks, Inc. Natick, MA).


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Model parameter, L for the permeation-only limit, was fit using Mathematica 4.1.5 (Wolfram Research, Inc., Champaign, IL).

Experimental Results Figure 1 depicts the resulting “release map” with respect to the sample composition. Test samples were exposed to 3 minutes of continuous 20 kHz ultrasound, and the final fraction of release was recorded as described above. Compositions examined and determined to be single phase by Veatch and Keller,42 the DAN-PC blue shift boundary found by Brown et al.,36 and the so-ld phase boundary published by Zhao et al.39 have been overlaid to assist interpretation in this non-raft-forming system. A general trend of reduced release with increasing cholesterol, consistent with observations from the raft-forming system in Small et al.,27 is apparent. Exceptions are noted as compositions exceed 0.6 xDPPC. These samples fall across the so-ld phase boundary from,39 thus these samples are no longer purely liquid phase. In similar systems51,52 so – lo coexistence and even 3 phase (ld-lo-so) coexistence has been observed in these regions of low cholesterol-high saturated lipid.


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Figure 1: POPC/DPPC/Cholesterol Release Map. Color scale is fraction of release after 180sec of continuous 20 kHz ultrasound exposure. □ samples denoted as single lipid-phase by Veatch and Keller 2003, ▬ ▬: DAN-PC blue shift boundary from Brown et al. 2007, ▬▬: so-ld phase boundary from Zhao et al. 2007.

The release map presents a mere “snap shot” of release. Establishing a thorough understanding of the effect of lipid phase behavior on release and discerning the possible mechanism(s) of release requires investigation of release kinetics. Accordingly, kinetic profiles for calcein release were obtained for all thirty-six compositions. In Figure 2 we examine the release profiles of samples with the same DPPC fraction, but varying ratios of cholesterol to POPC. The fraction of release is plotted against dimensionless time, Dt/a2, where D is the diffusion coefficient of the


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encapsulant in water, in this case calcein (3.5x10-6 cm2/s

53

), and a the (measured) diameter of

the vesicle. The use of this scaling parameter eliminates the effect of vesicle size (and when relevant, the type of encapsulated molecule) on the rate of diffusion within the vesicle aqueous core,54 meaning any variation in kinetic profiles are due solely to the different membrane permeabilities. It is important to note that using Dt/a2 does not eliminate the potential dependence of the effective permeability on the vesicle radius or encapsulant. In agreement with the release map in Figure 1, Figure 2 suggests that release decreases at high cholesterol content (greater than 0.45 x cholesterol). However the effect of cholesterol is non-monotonic, regardless of the POPC/DPPC ratio.


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Figure 2: POPC/DPPC/Cholesterol Dimensionless Kinetic Release by DPPC Mole Fraction. Panel (A) corresponds to 0.2 xDPPC and Panel (B) corresponds to 0.6 xDPPC. Cholesterol mole fractions are indicated in image inset. Fraction of release is ratio of release at time t to the total amount of calcein in the system (as measured using detergent dissolution). Dt/a2 is a dimensionless measure of time, where D is the diffusion coefficient of calcein in water (equal to 3.5x10-6 cm2/s [Prausnitz 1996]), and a the vesicle radius (measured via DLS).

Due to the three composition variables, it is important to examine entire release curves, thus in Figure 3, we plot POPC release profiles again, but arranged via cholesterol mole fraction. Panels A, B, C, and D are 0.15, 0.25, 0.35 and 0.50 x cholesterol respectively. Circles are 0.05 xDPPC, diamonds are 0.2 xDPPC, squares are 0.4 xDPPC, and triangles are 0.6 xDPPC. At all cholesterol fractions (those in Figure 3 and those not shown) the data appears to follow a single profile shape, indicating that samples with the same cholesterol mole fraction exhibit similar release kinetics. The average fraction of release near Dt/a2 = 5.0x106 for 0.15, 0.25, and 0.35 x cholesterol (Figure 3A, B, and C) are 0.64±0.07, 0.68±0.04, and 0.62±0.6 respectively. For 0.45, 0.50, and 0.55 x cholesterol (Figure 3D and data not shown) average fractions of release are 0.39±0.07, 0.43±0.04, and 0.46±0.5 respectively. This analysis suggests two distinct groupings of release rates; one for compositions near or below the DAN-PC blue shift boundary, and another for compositions of high cholesterol. There are two exceptions. In Figure 3B, 0.05 xDPPC shows a slower release profile than the rest, which may indicate a more cohesive membrane structure than samples near the DAN-PC blue shift boundary. The average fraction of release near Dt/a2 = 5.0x106 is 0.42; similar to the high cholesterol results. The membrane composition of 0.05 xDPPC and 0.25 x cholesterol is


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approximately 15% cholesterol below the DAN-PC blue shift. The other 3 DPPC compositions (0.2, 0.4, and 0.6) are within 5% cholesterol of the DAN-PC blue shift boundary found by Brown et al. Higher release near the DAN-PC transition may indicate perturbations in membrane packing due to the rearranging of the membrane from a liquid-disordered to liquid-ordered phase. When the membrane is fully liquid-ordered the release rates decrease as mentioned above. The second exception is Figure 3A the 0.80 xDPPC sample; it does not follow this single release shape. Phase maps indicate this sample may have solid-liquid coexistence. Data from Figure 3 suggests that the POPC/DPPC/Cholesterol system has a singular release rate when cohesive liquid phase is present. Any perturbations in the liquid phase (transition from disordered to ordered or presence of solid phase) cause release rates to increase.


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Figure 3: POPC/DPPC/Cholesterol Dimensionless Kinetic Release by Cholesterol Mole Fraction. Panel (A) corresponds to 15% cholesterol. Panels (B), (C), and (D) correspond with 25, 35, and 50% cholesterol respectively. ○ are 0.05 xDPPC, ◊ are 0.2 xDPPC, □ are 0.4 xDPPC, ∆


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are 0.6 xDPPC, and X are 0.8 xDPPC. Fraction of Release and Dt/a2 are as defined in Figure 2. Some so phase is present in the 0.6 and 0.8 xDPPC samples according to Zhao et al. 2007.

Derivation of a Two-Film Diffusion and Destruction Model Previously,27 we presented a model based on well-mixed compartment assumptions. Here we take a slightly different approach, which we believe more accurately describes the relevant physics, and present fits of our release profiles to a classic two-film system using Fick’s law for diffusion from a liposome consisting of aqueous core and lipid shell. Typical models assume that the concentration in the core varies slowly with time allowing us to apply the pseudo-steady state approximation. The fraction of encapsulated material, f, released by time, t, due to diffusion through the membrane can be modeled using a classic diffusion analysis to describe release from a sphere with a surface resistance:54,55

f p (t ) =

M (t ) =1− M∞ ∞

2

2

e − Dβ n t / a β n2 + L 2 − L 1 β n2 + L 2 − L

∑

β n2 (

)

∑

β n2 (

)

n =1 ∞ n =1

(1)

M∞ is the overall amount of encapsulated drug (or dye) in the system, and M(t) is the amount released by time t. D is the diffusion coefficient of the encapsulated material in water, and a is the vesicle radius. L is a dimensionless permeability parameter that defines the properties of the lipid bilayer resistance54,55 and can be defined as L=aDB/(hD), where h is the bilayer thickness, and DB is the effective diffusion coefficient through the bilayer.55 The eigenvalues βn are given by the roots of the expression:54


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β n cot β n + L − 1 = 0 ( 2 ) In the limit where L=0, the membrane is impermeable and no release takes place. We add destruction to this model by treating the rate of liposome destruction as an elementary, first-order reaction; that is, the rate of destruction of liposomes is directly proportional to the concentration of liposomes such that, in any given time period of US application, a fixed fraction of the vesicles is destroyed. Thus, the number (or concentration) of liposomes at any given time, N(t), is:

dN 1 = − N (3a ) τ dt N (t ) = N O exp(−t / τ ) (3b) where 1/τ is the characteristic rate of liposome destruction (analogous to a first-order reaction rate constant), and N0 is the initial number of vesicles. The fraction of release due to liposome destruction (assuming that all dye or drug is immediately released) is then given by

f d (t ) = 1 − exp[ − t / τ ] ( 4) Combining fp(t) with fd(t) gives a model for release via diffusion and vesicle destruction:

f tot = 1 − exp[ − t / τ ](1 − f p ) (5) where fp is as defined by equations (1) and (2). In the limit where no vesicle destruction takes place, τ→∞, and ftot = fp. In the opposite limit, where there is no diffusion, L and fp are zero, and ftot = fd. Model Results. We fit release data from POPC kinetic profiles in the two limits: When the membranes are impermeable (L=0) so that fp=0 and only destruction occurs or when τ→∞ so τ

that e-t/ =0 and only diffusion takes place. Figure 4 is a sample of these fits, where Figure 4A shows the destruction-only cases and Figure 4B shows the diffusion-only model. As can be seen


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both limits seem to fit the data equally well, suggesting that the release profiles, alone, cannot distinguish between the two mechanisms of diffusion and liposome destruction. In fact, POPC values for L and 1/τ are plotted against one another in Figure 5 and the relationship between the two is clearly linear.

Figure 4: Two-Film POPC/DPPC/Cholesterol Fits. Fitting LFUS-induced release assuming only vesicle destruction (A) or only diffusion (B). All vesicles contain 0.2 xDPPC and cholesterol as noted. Fit parameters for (A) are 1/τ = 1.70x10-7, 1.17x10-7, and 1.00x10-7 for cholesterol: 0.15(green), 0.45(red) and 0.6(blue), respectively. Fit parameters for (B) are L=5.08x10-7, 7.01x10-7 and 3.01x10-7, respectively.


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Figure 5: Relation of L to 1/τ. Fitted parameter L (from diffusion-only) in relation to 1/τ (from destruction-only) from the same kinetic profile. ○ are 0.05 xDPPC, ◊ are 0.2 xDPPC, □ are 0.4 xDPPC, ∆ are 0.6 xDPPC, and X are 0.8 xDPPC.

Experimental work by this group27 and others16,56 suggests vesicle destruction is a plausible mechanism of release but usually only a partial contributor. Simulations by Leontiadou et al.50 further support the possibility of vesicle destruction contributing to overall release. As in our previous work27 dynamic light scattering (DLS) was used to determine the presence of a destruction mechanism. Given that light scattering intensity is proportional to the sixth power of diameter and directly proportional to the number of scatterers,57 a decrease in intensity indicates smaller or fewer liposomes, or both. To differentiate between vesicle resizing and actual vesicle destruction, effective diameter was also measured to determine if any change in liposome size occurred. A decrease in scattering intensity, concomitant with a decrease in effective diameter would indicate vesicle resizing.57 Measurements were made on select samples to test for ultrasound-induced destruction. Values for scattering intensity and effective diameter are normalized by their initial measurements, prior to ultrasound exposure, to allow for clear comparison between samples from different preparation batches.


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Figure 6A indicated the normalized effective diameter of the samples according the DLS. Figure 6B shows the normalized scattering intensity. Red shapes indicate the sample composition was below the DAN-PC blue shift, and thus we expected them to behave like liquiddisordered samples. Blue shapes indicate sample compositions above the DAN-PC blue shift, and thus are expected to be liquid-ordered like. At all compositions, effective diameter showed minimal increase, on average a 35% increase. Increased vesicle size should result in increased scattering intensity if the vesicles were simply aggregating. However, scattering intensity decreased by 20% on average. This mild increase in size, concomitant with a small decrease in scattering intensity, suggests the destruction of some vesicles and the resulting formation of lipid debris

aggregates.

However,

unlike

the

liquid-disordered

samples

from

the

DOPC/DPPC/Cholesterol system studied previously27, in which average decrease in scattering intensity was 50% concurrent with a 162% increase effective diameter, destruction does not appear to contribute as greatly to release in this system. Nevertheless, the results of Figure 6 do indicate that LFUS-induced release might be explained – at least in part – by vesicle destruction.


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Figure 6: DLS destruction assay of select POPC/DPPC/Cholesterol compositions. Normalized effective diameter (A) and average count rate (B) from dynamic light scattering as a function of LFUS time (seconds). ♦ are 0.05 xDPPC, ▲are 0.2 xDPPC, and ■ are 0.4 xDPPC. Red points are those below the DAN-PC blue shift transition from Brown et al. 2007, and are expected to be liquid-disordered. Blue points are those above the transition line, and expected to be liquidordered.

In Figure 7 we plot the permeability (as fitted by equation 1) as a function of the cholesterol mole fraction; we distinguish between mixtures that are expected to be in the liquid-ordered phase (blue circle) and liquid-disordered phase (red diamond) based on the Brown et al.36 measurements, and those that may be in the two-phase disordered-solid coexistence region (green square) based on the Zhao et al.39 phase diagram.


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Perhaps surprisingly, we do not find a consistent correlation between the rate of release and the liquid phase type or cholesterol mole fraction; samples in the expected liquid-ordered regime did not display release rates (and thus, susceptibility to LFUS) that were consistently lower than those of samples in the expected disordered state. Moreover, Figure 7A clearly indicates that samples with the same cholesterol do not give identical values of L. However, there is clear evidence of two groupings of release rates in the single-liquid phase regime, in agreement with kinetic profile analysis from Figure 3. In samples with low cholesterol, average L for 15% cholesterol is 5.96 ± 1.56 x 10-7, 5.43 ± 1.79 x 10-7at 25% cholesterol, and 4.64 ± 2.02 x 10-7 at 35% cholesterol. For high cholesterol fractions L was 2.98±0.72 x10-7 at 45% cholesterol, 3.19 ± 0.47 x10-7 at 50% cholesterol, and 3.71 ± 0.69 x10-7 at 55% cholesterol. The most rapid release rates, and therefore most sensitive response to LFUS, occurred in the presence of ld-so phase coexistence. For example, at 15% cholesterol and 0.4 xDPPC, in the single liquid-low cholesterol regime, L was 7.76 x 10-7, but over three times greater, 2.34x10-6, at 0.8 xDPPC where so phase is present. These results suggest that for these types of lipid mixtures, LFUS response is not sensitive to the type of liquid phase, but rather to the presence of domain boundaries between distinctly differing phases.


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Figure 7: The effect of composition on the rate of release, as defined by the effective permeability L (equation 1). (A): L as a function of cholesterol mole fraction, x. ● denote cholesterol values above the transition line measured by Brown et al. 2007, and are therefore expected to be in the liquid-ordered phase. ♦ are below those values, and are therefore expected to be in the disordered phase. ▲ are those compositions that fall on the transition line. ■ denote mixtures that are in the potential disordered-gel two-phase regime, as measured by Zhao et al. 2007. (B): L as a function of the ternary composition. □ indicates compositions of a single liquid phase by Veatch and Keller 2003. ▬ ▬ is interpolated DAN-PC blue shift boundary from Brown et al. 2007. ▬ is so-ld boundary redrawn from Zhao et al. 2007. Color bar indicates value of dimensionless diffusion-only fit parameter L. Discussion and Conclusions In this paper we investigate the effect of bilayer composition on LFUS-induced release from POPC/DPPC/cholesterol vesicles. Bilayer sensitivity to LFUS was measured through the rate of release of a self-quenching fluorescent molecule from liposomes; bilayers that are more resistant to LFUS are expected to exhibit longer release times, and slower release rates, than ones that are more sensitive and thus easily disrupted.


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Previous studies have shown that the rigidity and moduli of lipid bilayers are highly sensitive to the cholesterol content.58-61 Thus, it may have been expected that the rate of LFUS-induced release would decrease systematically with increasing cholesterol mole fraction. Yet, as shown in Figures 1, 2, and 7 , both the extent and the rate of release do not vary systematically for systems with a fixed DPPC mole fraction, nor due bilayers with the same cholesterol mole fraction give similar release rates. However, data indicates two groupings of release rates, dependent on the cholesterol mole fraction, seen most clearly in Figures 3 and 7A. The only exceptions are samples where a solid phase is expected to be present. Two mechanisms have been suggested for LFUS-induced release from lipid bilayers: Vesicle destruction and increased bilayer permeability (namely, diffusion).27 As shown in Figure 4, the release profiles can be fit equally well by both mechanisms (Figure 4). Thus, distinguishing between the two mechanisms requires direct examination of the two mechanisms. To that end, we performed DLS-based destruction assays on the POPC/DPPC/Cholesterol vesicles. As shown in Figure 6, only minor increases in effective diameter, and similarly small decreases in scatter intensity were observed, suggesting that destruction does not contribute greatly to release in this system, and that release is dominated by bilayer permeation. Previous work on DOPC/DPPC/cholesterol bilayers suggests that the sensitivity of bilayers to LFUS depends on the lipid phase; bilayers in the liquid-ordered phase, where the lipids are closely packed, were found to be less sensitive to LFUS than liquid-disordered phases.27 In the coexistence two-phase regime the sensitivity to LFUS decreases with increasing fraction of the ordered phase.27 However, the phase diagram of POPC/DPPC/cholesterol differs significantly from that of the DOPC system, so that direct comparisons cannot be made. Throughout the greater part of composition space, the POPC bilayer is in one liquid phase.37,42 Brown et al.36


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found that a fluorescent probe that partitions preferentially to the liquid-disordered phase displays a shift in the emission wavelength, indicating a change in environment, which was taken to correlate with system transition into the single phase liquid-ordered regime. More recently, Zhao, et al.39 suggest that there may be a region of coexisting ld -so phases. In Figures 7 A and B we plot the permeability as a function of the cholesterol mole fraction and as a function of membrane composition. A low value of L indicates low rate of transport, namely, weak sensitivity to LFUS, while a high L value corresponds to rapid release and high LFUS sensitivity. We did not find a consistent correlation between the rate of release and the liquid phase type. However, the presence of domain boundaries between distinctly differing phases of liquid and solid is found to cause release rates to more than double. The line tension between so and ld phases is expected to be high due to the (relatively) large thickness mismatch, and the rigidity of the so phase.62,63 Thus, the contact line between phases is likely to ‘fail’ under the LFUS-induced pressure. In contrast, the line tension between liquid domains (ld and lo) is appreciably smaller, since the thickness mismatch is smaller and both phases can deform around the contact line.24,62-66 Thus, in the DOPC system the LFUS-induced perturbation is less likely to affect bilayer integrity in regions of coexistence when compared to the POPC system. We thus interpret the results of this as evidence that membranes with large thickness mismatches, such as ternary systems of unsaturated lipids, cholesterol, and sphingomyelins51,52 or cerebrosides,67 may be sensitive to LFUS even in the liquid-liquid coexistence regions. To further confirm the validity of the permeation model, we compare the values of L to expected, literature-based values: The permeability L is defined, in the two-film model, as aDB/(hD), where h is the bilayer thickness, and DB is the effective diffusion coefficient through the bilayer.55 Using bilayer thickness of h = 4.32nm for our system,68 D value for calcein


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diffusivity in water of 3.5x10-6 cm2/s,53 and the a values determined from DLS yield that the diffusion coefficient through the membrane (due to LFUS) DB should range from 3.45 x 10-14 to 2.95 x 10-13 cm2/s. There are no literature values for LFUS-induced effective diffusivities of calcein across lipid-bilayers although gap junctions in C6 cells that transmit calcein across a lipid monolayer are known to have an effective diffusivity of 2.5 x 10-9 cm2/s.69 Considering the fact that the LFUS field permeates the bilayer only a fraction of the time, the values we find for L seem reasonable. In conclusion, we examined the effect of POPC/DPPC/cholesterol bilayer composition on membrane sensitivity to LFUS by monitoring LFUS-induced release from unilamellar vesicles. The release profiles are fit equally well by a liposome destruction model and a bilayer permeation model, thus the mechanism of release cannot be determined by this model alone, emphasizing the importance of destruction assays. These suggest that permeation is the dominant release mechanism, and the values obtained for the bilayer permeability are in agreement with expectations for this system. Generally, we find that the sensitivity of the POPC bilayers to LFUS (as measured by the rate of release) increases with increasing DPPC content, as the bilayer tends toward solid-ordered. Sensitivity decreases with either POPC or cholesterol mole fractions, which “melt” the bilayer to a more disordered state by disrupting packing.59,70 In the liquid regime of this system there is no recorded phase transition. However, compositions near the DAN-PC blue shift boundary are more sensitive to LFUS than samples with high cholesterol, thus cholesterol is the determining factor in release rates. In our previous paper27 we determined that phase coexistence did not enhance LFUS release in bilayers with liquid-ordered/liquid disordered coexistence; however


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data from this study indicates that solid-liquid phase coexistence does enhance membrane sensitivity to LFUS.

Acknowledgements The authors would like to thank Chris Bawiec for measurement of ultrasound field parameters. This work was supported in part by the National Science Foundation under Grant number DGE0947936.

AUTHOR INFORMATION Corresponding Author *Email: [email protected], Tel: 1-215-895- 6694, Fax: 1-215-895-5837 Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Funding Sources This work was supported in part by the National Science Foundation under Grant number DGE0947936. ACKNOWLEDGMENT The authors would like to thank Chris Bawiec for measurement of ultrasound field parameters. This work was supported in part by the National Science Foundation under Grant number DGE0947936.


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ABBREVIATIONS DAN-PC, 1-myristoyl-2-[12-[(5-dimethylamino-1-naphthalenesulfonyl)amino]dodecanoyl]-snglycero-3-phosphocholine; DANSYL, 5-(dimethylamino)naphthalene-1-sulfonyl; DLS, dynamic light scattering; DOPC, 1,2-dioleoyl-sn-glycero-3-phosphocholine; DPPC, 1,2-dipalmitoyl-snglycero-3-phosphocholine; EDTA, ethylenediaminetetra-acetic acid anhydrous; kcps, kilocounts per second; ld, liquid-disordered; LFUS, low frequency ultrasound; lo, liquid-ordered; LUV, large unilamellar vesicles; MLV, multilamellar vesicles; NaCl, Sodium Chloride; POPC, 1-palmitoyl2-oleoyl-sn-glycero-3-phosphatidylcholine; so, solid-ordered; TrisBase, Trimethylol Aminomethane; US, Ultrasound.

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Low-Frequency Ultrasound-Induced Transport across Non-Raft

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