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Applied Acoustics xxx (2008) xxx–xxx
Contents lists available at ScienceDirect
Applied Acoustics
journal homepage: www.elsevier.com/locate/apacoust
Phospholipid-stabilized microbubbles: Influence of shell chemistry
on cavitation threshold and binding to giant uni-lamellar vesicles
Steven P. Wrenn a,*, Michał Mleczko b, Georg Schmitz b
a
b
Department of Chemical and Biological Engineering, Drexel University, 3141 Chestnut Street, Philadelphia, PA 19104, United States
Lehrsthul for Medizintechnik, Ruhr University Bochum, Universitätsstraße 150, 44780 Bochum, Germany
a r t i c l e
i n f o
Available online xxxx
Keywords:
Microbubbles
Cavitation
Vesicles
Avidin
Biotin
Phospholipid gel
a b s t r a c t
We demonstrate the feasibility of covalently linking a single microbubble to a single, giant uni-lamellar
vesicle (GUV). Such a combination of GUV plus microbubble might prove useful as a new drug delivery
vehicle involving microbubble cavitation-induced sonoporation of the vesicle bilayer as a release mechanism. We therefore applied the well known methodology of passive cavitation detection to measure the
influence of lipid shell chemistry on inertial cavitation thresholds for externally added microbubbles. We
find that cavitation threshold changes significantly with changes in either molecular weight or mole fraction of poly(ethylene glycol), historically used to impede gas dissolution and microbubble coalescence.
We attribute changes in cavitation threshold to changes in microbubble resonance frequency resulting
from changes in microbubble shell bending elasticity. To further demonstrate the influence of shell chemistry on microbubble behavior, we describe how several common bubble phenomena – and some new –
respond to changes in lipid chain length.
Ó 2008 Elsevier Ltd. All rights reserved.
1. Introduction
Ultrasound contrast agents, which are microbubbles used for
enhancement of ultrasonic images, have gone through several generations of development [1]. The common feature is a gas core plus
a stabilizing shell. First-generation agents comprised air plus a
shell of albumin, lipid, or acrylate.1 Second-generation agents improved upon the first-generation by employing gases other than
air, typically fluorinated compounds (octafluoropropane, perfluorobutane, or sulfur hexafluoride). Owing to lower solubility in water
and slower diffusivity – relative to air – microbubbles produced from
these gases were more long-lived than their first-generation counterparts. Third-generation agents built upon the second-generation
agents by incorporation of species into the stabilizing shell. The purpose of theses species is typically to convey added stability (e.g.,
charged surfactants or PEGylated lipids to prevent bubble coalescence) or to enable targeting to a specific receptor within tissue
(via receptor ligands, analogous to avidin-biotin binding). More recent research activity involves design of microbubbles along similar
lines but for the purpose of targeted or controlled drug delivery
rather than for imaging [2–5].
It is this latter aspect of microbubble design and synthesis that
interests us. In particular, we are interested in designing a drug
* Corresponding author. Tel.: +1 215 895 6694; fax: +1 215 895 5837.
E-mail address: spw22@drexel.edu (S.P. Wrenn).
1
One can also speak of so-called generation zero contrast agents, which were
merely free air bubbles (and exceedingly unstable).
delivery vehicle that combines microbubbles with giant phospholipid vesicles. Others have combined microbubbles with small, unilamellar vesicles [4]. Our vehicle combines the advantages of
stealth liposomes (relatively large drug carrying capacity and ability to carry either hydrophobic or hydrophilic drugs) with the
acoustic activity of microbubbles (which enables use of ultrasound
as a remote trigger to stimulate drug release from vesicles via
sonoporation).
Earlier we reported release of fluorescent drug mimics from
phospholipid vesicles using low-frequency ultrasound [6].
Although no microbubbles were utilized in that work, we believe
the release mechanism necessarily involved cavitation of gas bubbles that developed from gas voids or dissolved gases contained
within the aqueous system (rather than a direct interaction between the vesicles and ultrasound wave). A question naturally
arises, namely what is the mechanism of cavitation responsible
for release?
As with sonoporation of cell membranes, there are two possibilities [7]. One, perhaps the most generally accepted, is so-called stable cavitation, whereby shear stresses associated with steady
streaming of fluid close to the phospholipid membrane are sufficient to tear the membrane [8–11]. A second is transient, or inertial, cavitation [12–14]. Here microbubbles expand sufficiently
during a single ultrasound cycle to collapse, literally, on themselves; one consequence of collapse is generation of a shock wave
capable of rupturing a nearby phospholipid membrane. The distinction between stable and inertial cavitation is conceptually
clear, and experimental determination of cavitation threshold (that
0003-682X/$ - see front matter Ó 2008 Elsevier Ltd. All rights reserved.
doi:10.1016/j.apacoust.2008.09.017
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is, the ultrasound intensity above which bubbles collapse violently
during a single ultrasound cycle) based on changes in acoustic
spectral emissions – or passive cavitation detection – is well described [15,16].
While the passive detection methodology–for liquids containing native bubbles – is well known, the application of passive
detection to liquids containing synthesized, externally added
microbubbles is not [17]. There are no experimental accounts of
how microbubble cavitation threshold correlates with microbubble lipid shell composition. Yet such information could be very useful, as lipid shell chemistry is a potential tuning parameter to
achieve – or avoid – cavitation in targeted drug delivery applications. This is because of how cavitation threshold relates to microbubble resonance frequency via microbubble shell bending
rigidity, which we summarize below.
Viewing a (native, spherical ideal gas) bubble – suspended in a
liquid – as a simple mechanical oscillator, its resonance frequency
is given as
1
xo ¼
Ro
sffiffiffiffiffiffiffiffiffiffiffi
3jPo
ð1Þ
q
where xo is the bubble (angular) resonance frequency, j is the
polytropic index, Po is the liquid hydrostatic pressure, and q is the
liquid density [7]. For a synthetic microbubble encapsulated by a
monolayer shell, de Jong et al. [18] have shown the resonance frequency to be
x2 ¼ x2o þ
Sshell
m
ð2Þ
where x is the (angular) resonance frequency of a shell-containing
microbubble, m is microbubble mass and Sshell is the so-called shell
parameter, which de Jong et al. show is proportional to shell stiffness. One can therefore express microbubble resonance frequency
in terms of measurable shell material properties familiar to colloid
scientists:
x2 ¼ x2o þ
8pkb
ð1 mÞmh
2
ð3Þ
where m is the Poisson ratio, h is monolayer shell thickness, and kb is
the membrane rigidity or bending elastic modulus. kb is directly
proportional to Gibbs elasticity (or area expansion modulus) of a
membrane, which is readily measured [19]. Relevant to this study,
poly(ethylene glycol) and lipid chain length are known to influence
membrane rigidity appreciably [20–25].
Knowing how a microbubble’s resonance frequency depends on
its size and shell bending elasticity, we now turn our attention to
inertial cavitation of the microbubble. In the limit of very large
bubble resonance frequency – or very small bubble size – there
is ample time during a single ultrasound cycle for the bubble to expand to its thermodynamic limit. This is known as the quasi-static
regime, and the cavitation threshold pressure becomes the familiar
Blake threshold [7].
On the other hand, when microbubble resonance frequency is
sufficiently small that quasi-static conditions no longer apply
(e.g., larger bubbles or bubbles with modified shells), then there
is insufficient time between successive negative peak pressures
to allow complete bubble expansion. The situation becomes exacerbated with increasing driving frequency and decreasing bubble
resonance frequency such that transient cavitation becomes
increasingly difficult to achieve as one increases sonication frequency. The effect is shown graphically by Apfel and Holland
[26] and reproduced by Leighton [7], who also describes the effect
mathematically by combining the works of Apfel [27] and Flynn
[28] to give
rffiffiffiffiffiffiffi
1=3 pffiffiffiffiffi
3
q
2
Pt
Pt ¼ P o þ ðRt xÞ
ðPt Po Þ
1þ
2
3P o
2j
ð4Þ
where Pt is the cavitation pressure threshold, Rt is the bubble
threshold radius, above which cavitation is expected, Po is the original hydrostatic pressure in the liquid outside the bubble, j is the
polytropic index, q is the liquid density, and x is the frequency of
the sound wave.
Eq. (4) shows how cavitation threshold is sensitive to microbubble resonance frequency. Coupling this result with Eq. (3) makes
clear that a change in microbubble shell chemistry, via its influence
on shell bending rigidity and therefore microbubble resonance frequency, can be expected to impact cavitation threshold. We seek to
demonstrate this expected effect experimentally.
The purpose of this study is threefold: (1) to demonstrate feasibility of linking, via avidin and biotin, a giant uni-lamellar vesicle
(GUV) to a third-generation microbubble and (2) to measure the
influence of microbubble lipid shell chemistry on cavitation
thresholds. The first task is necessary because we ultimately intend
to use microbubble cavitation to induce sonoporation of the vesicle
bilayer; this requires that a vesicle and microbubble be in close
proximity (closer than can be achieved with reasonable particle
concentrations). The second task complements the first task because knowing how microbubble composition impacts cavitation
threshold will aid in the design of a delivery vehicle whose cavitation pressure threshold can be set to a desired value (3) beyond
these two specific goals, a more general – but key – point of this
work is to demonstrate that a microbubble shell offers more benefits than a mere barrier against gas diffusion and an impediment
against microbubble coalescence. Although our primary interest is
in microbubble shell chemistry as a potential tuning parameter
that will allow control over cavitation – or lack thereof – in drug
delivery applications, we wish to draw attention to the importance
of shell chemistry on a myriad of bubble phenomena. We therefore
begin with a review of commonplace microbubble synthesis,
showing how simple changes in lipid chain length give drastically
different microbubble behavior.
2. Materials and methods
2.1. Microbubble synthesis
Microbubbles were made according to previous reports [29,30].
Specifically, distearylphosphatidylcholine (DSPC, a free sample
from Lipoid GmbH, Ludwigshafen, Germany) and polyethyleneglycol6000monostearate (PEG6000MS, a free sample from Stepan,
Inc.) were combined in chloroform so as to give a mixture comprising 95 mol% DSPC. Chloroform was removed under a stream of
nitrogen, and the resulting dried film was hydrated in a buffer
solution containing 0.9 wt.% NaCL and 5 mM HEPES by direct sonication for 30 s (at 24 kHz, using a Hielscher UP400S sonicator with
a 7 mm tip at the lowest power setting). The aqueous suspension,
comprising 2 mg/mL DSPC and 1 mg/mL PEG6000MS, was then
placed in an 80 °C oven for 1 h (to ensure adequate lipid mixing
and hydration with the lipids in a liquid state).
Microbubbles were prepared by dispensing 6 mL of the aqueous
lipid suspension of multi-lamellar liposomes into a 20 mL scintillation vial and positioning the vial under the same sonication tip
used to hydrate the lipid film. The sonicator tip was submerged
1 cm beneath the liquid surface. Also beneath the liquid surface,
and directly beneath the sonicator tip (the positioning here is
important; the gas cannot be sparged merely anywhere within
the liquid but must be delivered within the sound field), was a syringe needle connected to a flexible tube, which was in turn connected to a gas cylinder containing sulfur hexafluoride (SF6).
Fig. 1 is a schematic drawing of the actual set-up. SF6 was delivered
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electrical
connection
sonicator
tip
electrodes
gas delivery
needle
electrical
connection
Fig. 2. GUV Synthesis chamber: giant uni-lamellar vesicles (GUVs) are made by
electroformation in a housing comprising four indium tin oxide-coated microscope
slides (two on top, two on bottoms). A lipid film is deposited onto the bottom two
slides and dried, the bottom chamber is filled with a glucose solution, the top
chamber is pressed into the bottom chamber to give a water-tight seal, and
(alternating) current is delivered (see text for details).
Fig. 1. Microbubble synthesis by sonication: microbubbles are prepared by direct
sonication (24 kHz, Hielscher UP400S) while sparging SF6 into an aqueous
suspension of phospholipid multi-lameller vesicles. Gas is sparged through a
syringe needle located directly beneath the tapered sonicator probe.
a box with an internal volume of 10 mL. Holes were drilled, and
copper leads soldered onto the ITO slides, such that filling the
chamber with water would give a complete electrical circuit.
Prior to filling the chamber with water, 1 mL of a solution comprising 10 mg/mL Egg PC (in chloroform) plus 50 lL of 50 mg/mL
PEG6000MS (in chloroform) was deposited onto the two ITO slides
on the bottom, female half of the chamber and allowed to dry completely under a stream of nitrogen. After drying, the female chamber was filled with 12 mL of a 50 mM glucose solution; the purpose
of glucose was to provide optical contrast during light microscopy.
The male half was then pressed into the female half, causing 2 mL
water to spill out (this guaranteed no gas voids within the chamber
and gave a water-tight seal). A sinusoidal voltage (amplitude 6 V
peak-to-peak, frequency 10 Hz) was applied using a programmable
signal generator (Agilent 33120A, Santa Clara, CA) for one and a
half hours, followed by harvesting of glucose-filled GUVs (of nominally 1–10 lm diameter).
at a rate of nominally 1 mL per minute (this was not measured but
was estimated from the rate and size of (macro) bubble formation
at the needle tip in the absence of sonication).
During gas delivery, direct sonication was applied at maximal
power for 30 s. Immediately after sonication, the sample was removed from the sonicator assembly and quenched in cold water
to ensure the lipids were in the gel phase. Sample appearance
changed from a cloudy liquid into a bright white foam after several
seconds of sonication and separated upon standing into three distinct phases; an upper layer of macroscopic bubbles, a lower aqueous (clear) layer, and a middle layer containing microbubbles (this
layer appeared milky and became narrower with time as bubbles
floated upward; that is, the upper phase boundary remained stationary, but the lower boundary rose with time).
It should be stated that the above formulation and procedure
gave the best results in our hands. Time and intensity of sonication,
fraction of PEG6000MS, and lipid type (replacing DSPC with dimyrstoylphosphatidylcholine – DMPC, dipalmitoylphosphatidylcholine – DPPC, and Egg PC – which comprises a mixture of
phosphatidylcholines with varying chain lengths and degrees of
chain unsaturation) were varied but gave inferior results. Nevertheless, these variations led to interesting observations, described
herein.
Biotinylated-PEGylated-distearoylphosphatidylethanolamine
(Biot-DSPE) was purchased from Avanti Polar Lipids (Alabaster,
AL), and native avidin was purchased from Affiland (Liège, Belgium). Separate populations of biotinylated bubbles and biotinylated GUVs were prepared by including 1 mol% Biot-DSPE in each of
the formulation recipes. A stock solution of 5 lg/mL Avidin was
also prepared separately. In a typical binding experiment, 20 lL
of biotinylated bubbles were added (immediately after preparation
via sonication) to 500 lL of biotinylated GUVs, followed by addition of 20 lL of Avidin and 500 lL of 50 mM sucrose solution with
gentle mixing after each addition.
2.2. Giant uni-lamellar vesicle (GUV) preparation
2.4. Imaging system
GUVs were prepared by a slight modification of methods described previously [31,32]. Briefly, a home-built electroformation
chamber was constructed, in which four indium-tin-oxide (ITO)coated glass slides were glued to the inside of two halves (two
slides on the top, male half, and two on the bottom, female half)
of a plastic housing (see Fig. 2). The male and female halves of
the housing – when mated together with rubber gaskets – formed
Bubbles and GUVs were observed with a system described previously [33–35]. Briefly, samples were inserted via syringe into a
200 lm inner diameter (8 lm wall thickness) CUPROPHANÒ
RC55 cellulose capillary (Membrana GmbH, Wuppertal, Germany)
that was isolated within a de-ionized water-filled Perspex tank,
illuminated from below with a DC 421 40,000 footcandle fiber optic continuous light source (Stockeryale, Inc., Salem, NH), and
2.3. Avidin-biotin binding
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observed from above through a LUMPlan FL 60 high numerical
aperature (NA = 0.90) water immersion objective lens (Olympus
Deutschland GmbH, Hamburg, Germany). The Perspex container itself was mounted on top of a custom-built micron-adjustable x–y
table, and the microscope was connected to a video camera. A
spherically focused, 2.25 MHz (Panametrics, Inc., Waltham, MA)
transducer was oriented perpendicular to the capillary and at 90°
relative to both the light source and the microscope objective.
The configuration was such that optical focal and acoustic focal
regions overlapped, and the capillary was positioned within the
region of overlap. Fig. 3 shows an image of a typical batch of
DSPC-stabilized microbubbles photographed within the cellulose
capillary.
2.5. Cavitation detection system
A custom passive cavitation system was designed and built,
based on well established protocols [15–17]. Fig. 4 provides a
schematic drawing of the physical layout of the system. Two
Olympus-NDT (Waltham, MA) V395 focused ultrasound transducers, each with a 2.25 MHz center frequency, are oriented at exactly 90° relative to one another within a plexi-glass, de-ionized
water-filled tank. One transducer acts an excitation source and
transmits pulsed ultrasound at a single frequency. A second transducer serves as an emission detector and operates over a wide
range of frequencies (so that it can detect harmonics, sub-harmonics, and broadband noise). As with the microbubble imaging
system, the container is outfitted to house a light source and
microscope so as to allow simultaneous optical and acoustic
imaging.
The region of focal overlap between transducers is 1 mm3, and
the bubble concentration within the detection system is diluted
until no more than one bubble is observed within the focal region
at any given time (which requires that in some cases no bubble is
observed). While within the focal region, a bubble experiences
multiple excitation pulses, and bubble response to each pulse is
measured and recorded. Hundreds of bubbles are sampled in this
same manner at a given power setting, and the power is then adjusted – and the process repeated – so as to determine bubble response over a range of acoustic intensities.
Fig. 3. A typical batch of microbubbles: shown are microbubbles comprising an SF6
core and a shell of 95 mol% DSPC plus 5 mol% PEG6000MS. DSPC concentration was
2 mg/mL in the original formulation, which yields on the order of 109 bubbles per
mL.
Bubble excitation was achieved in two different ways so as
to give two versions of the detection system shown in Fig. 4.
In the first version, the system was operated so as to generate
acoustic spectrograms. Ultrasound excitation consisted of a
10-cycle pulse with a center frequency of f0 = 1.8 MHz. The
insonification pulses were generated by a programmable pulser/receiver system (Inoson PCM100, Inoson GmbH, St. Ingbert,
Germany). The pulser/receiver operated in transmission mode,
and the spectrum of each scattered signal was determined and
plotted as a spectrogram.
Acoustic spectrograms are appealing in that they provide dramatic visual evidence – or lack thereof – of inertial cavitation at
any particular acoustic intensity. However, they do not lend themselves well to examining a wide range of intensities simultaneously. Accordingly, a second version of the cavitation detection
system was achieved by using an arbitrary waveform generator
(Agilent 33250 A, Agilent Technologies Inc., Santa Clara, CA) to
transmit a 5-cycle sine burst with a center frequency of
f0 = 2.25 MHz. Another arbitrary waveform generator (Agilent
33120 A, Agilent Technologies Inc., Santa Clara, CA) was used to enable a triggering of 50 bursts with a repetition frequency of 250 Hz.
The repetition frequency of the pulse trains was 1.4 Hz (see
Fig. 3b). Scattered signals were amplified using an Olympus-NDT
5910R pulser/receiver operated in passive mode. The amplified signals were recorded for further off-line processing using an Acqiris
DP-310 digitizer card. A matched filter, set-up for the detection of a
five-cycle sine burst at f0 = 2.25 MHz, was used to detect whether a
bubble was present in the acoustical focus. If detection failed for
the following burst, the bubble was counted as destroyed. Otherwise it was counted as survived.
3. Results and discussion
3.1. Bubble synthesis and characterization
A key aspect of bubble synthesis relates to the nature of the
shell [36,37]. We favor phospholipids, which are used in commercial formulations [1,38], are commonplace in academic formulations [39–42], and – as they are components of native
cell membranes – are inherently biocompatible. Moreover,
phospholipid vesicles serve as cell membrane mimics, and study
of phospholipid bilayers and associated phase behavior is a mature field within the biophysics community [43–48]. We are not
the first to appreciate this, as others have already demonstrated
analogies between phospholipid bilayer systems and bubble
phospholipid monolayers; particularly exciting among these
are systems demonstrating multi-phase coexistence and examinations of cholesterol [49–51]. Such studies are not only remarkable scientifically but might prove useful to the extent that
cholesterol-rich domains, found in cell membranes and known
as ‘‘lipid rafts” [52–54], play a role in sonoporation (we anticipate that rafts might influence cell membrane susceptibility to
rupture).
Of particular importance to microbubble synthesis, the chain
melting temperature of a phospholipid (or gel transition temperature, i.e., the temperature at which the phospholipid changes between a liquid and a gel phase) depends in a known way on
chain length (that is, the number of methylene units) and chain
unsaturation (that is, the number of carbon–carbon double bonds)
[19,55]. If one examines the microbubble literature, one finds DSPC
is ubiquitous in formulations based on phoshpholipids. However,
there is nothing special about DSPC per se; it just so happens to exist as a gel phase at body temperature. One could just as easily use
DMPC, if the temperature of application were below 25 °C. Moreover, DSPC becomes ineffective as a shell material at temperatures
above 60 °C. We demonstrate these effects here, stressing the point
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Fig. 4. Cavitation detection system: (a) the cavitation detection system consists of two spherically focused ultrasound transducers, one utilized for transmission and a second
– at a 90° relative to the first – used to receive acoustic waves, across a range of frequencies, re-emitted by bubbles within the region of focal overlap. Raw acoustic data is
used to generate acoustic spectrograms, and inertial cavitation events are detected as a sudden appearance of broadband noise; (b) alternatively, one can test for cavitation as
the disappearance of a bubble upon successive excitations using a pulse train to excite bubble samples.
that synthesis of stable microbubbles requires that a phospholipid
be in the gel phase.
Table 1 lists three phospholipid species, along with their chain
melting temperatures, and our success – or lack thereof – in making stable microbubbles with these species at a few different temperatures. For all cases in which the processing temperature was
below the chain melting temperature, irrespective of the phospholipid identity, bubble synthesis was feasible. Conversely, bubble
synthesis at temperatures greater than the chain melting temperature was not feasible, regardless of which phospholipid was used.
It is clear that having a gel-phase lipid is the key to preventing gas
escape.
Table 1
Influence of gel transition temperature on microbubble synthesis.
Phospholipid species
Gel transition temperatures (°C)
7 °C
37 °C
60 °C
Egg PC
DMPC
DSPC
10
23
56
Noa
Yes
Yes
No
Noa
Yes
No
No
No
a
In some cases bubbles survived at these temperatures, having been prepared
initially at cooler temperatures. However, the bubbles which survived grew
uncontrollably to fill the capillary. It was not determined unequivocally whether
growth resulted from simple volume expansion of individual bubbles or if diffusion
played a role, but growth did not appear to result from coalescence or fusion of
multiple bubbles (Note: All bubble formulations included 5 mol% PEG6000MS).
An interesting observation, not evident from Table 1 is what
happens when bubbles – made initially with phospholipids in a
gel phase – are heated above the phospholipid chain melting temperature. We find that such bubbles swell significantly, from an
initial diameter of several microns to a final size that is hundreds
of microns. Indeed, these bubbles expanded so as to fill the capillary in which they were contained and therefore became nonspherical, taking on the geometry of the capillary itself. This is
shown in Fig. 5. In particular, panels g–i show how bubbles made
from DMPC at 7 °C grow spontaneously upon heating to 37 °C. Panel f shows how similar growth occurs for Egg PC at 7 °C. No such
growth is observed for DSPC, which remains in the gel phase at
these temperatures (Note: imaging of DSPC at a temperature above
its chain melting temperature was not possible because of equipment sensitivity). It was not determined unequivocally whether
growth resulted from simple expansion of individual bubbles or
if inward diffusion of gas was involved; growth did not appear to
involve coalescence or fusion of multiple bubbles.
Another interesting observation, described at least once previously [56], is a cyclical process whereby microbubbles are believed
to shed excess phospholipids as they lose gas to diffusion and dissolution. This is shown in Fig. 6, which gives images of the actual
process and a cartoon representation describing what is believed
to be occurring. Although no single bubble is ever perfectly spherical, the approximate geometry of a bubble is that of a sphere.
Moreover, the gas–liquid interface at the bubble perimeter usually
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Fig. 5. Influence of phospholipid chain length and phase (Gel versus liquid): (a) DMPC at 7 °C (magnified view – stable bubbles), (b) DSPC at 37 °C (magnified view – stable
bubbles), (c) Egg PC at 7 °C (no bubbles), (d) DMPC at 37 °C (no bubbles), (e) DSPC at 37 °C (stable bubbles, observed here as dark band of bubble population), (f) Egg PC at 7 °C
(in some cases, growth), (g–i) DMPC initially prepared at 7 °C, then heated to 37 °C, (panels g–i show 10 min time sequence of bubble growth, which appears to be a
combination of expansion or growth of individual bubbles – or both – rather than coalescence or fusion of multiple bubbles). Notes: (1) Phospholipid chain melting
temperatures are 10 °C, 23 °C, and 56 °C, for Egg PC, DMPC, and DSPC, respectively. (2) Capillary diameter is 200 lm in each of panels c–i, and magnification in panels a and b
is 10 that of panels g. (3) All bubble formulations included 5 mol% PEG6000MS.
appears smooth. As gas inevitably diffuses through the shell wall
and dissolves into the bulk liquid phase2 a bubble temporarily loses
its spherical geometry and smooth surface and appears to crumple
or buckle (Fig. 6, top left) as phospholipids attempt to conserve area
(think of a helium balloon a few days after the party). As the gas core
continues to shrink, the excess curvature associated with crumpling
becomes excessive such that shedding of a phospholipid aggregate
becomes favorable.
Borden and Longo refer to this as the ‘‘zippering” effect [56]
whereby apposing portions of the crumpling monolayer shell contact one another to form a bilayer. Given the trade-off in energies
for forming a new bilayer fragment versus creating higher curvature, they also found the zippering process depends on phospholipid chain length (for example, short chain phospholipids
exhibited continual bubble shrinkage without obvious crumpling,
suggesting constant shedding of lipid). We would like to add the
observation that incorporation of biotinylated lipids within the
bubble shell also affected bubble geometry, leading to ‘‘cigarshaped” structures upon dissolution (and this effect was more pronounced in the presence of an ultrasound field).
For bubbles which exhibit crumpling, the nascent bilayer fragment is seemingly ejected from the bubble shell into the liquid
phase – where it most likely folds on itself to eliminate aqueous
exposure of hydrophobic phospholipid chains at the edges and
2
This is true even if the liquid is initially saturated with gas, as described in 1950
by the seminal work of Epstein and Plesset [66]. Note; the original paper by Plesset
appears to contain an algebraic error as one attempts to derive Eq. 34 from Eq. 31.
Both equations are correct as written. The seeming contradiction stems from the fact
that the parameter Cs represents bulk phase saturation in Eq. 31 and bulk phase
saturation in the presence of a bubble in Eq. 34. The point is that the presence of a
bubble (and associated interfacial tension and curvature) changes the saturation
value. For an excellent discussion of the topic (see the work of Duncan and Needham
[67]).
thereby forms a small, uni-lamellar vesicle. The shedding of the bilayer fragment causes the bubble to revert or ‘‘snap back” to a
mostly spherical geometry with a smooth interface (Fig. 6, top
right). The process then repeats in a cyclical fashion (Fig. 6,
bottom).
In addition to phospholipid species, it is worth noting that bubble behavior and stability also appear to depend somewhat on processing conditions. Although difficult to quantify, we found that
higher sonication intensities during gas delivery led to higher bubble yields and more stable (that is, longer-lasting or more diffusion-resistant) bubble populations. This was not always the case,
suggesting other parameters are at play, but was a generally observed feature. We believe this is easily explained by tighter lipid
chain packing resulting from higher peak pressures at higher sonication powers.
A final point to make concerning bubble synthesis and behavior
relates to the incorporation of PEGylated lipids within the monolayer shell. Just as it is essential to have a gel-phase phospholipid
to prevent excessive gas escape from a single bubble, it is likewise
necessary to have a shell component that prevents excessive coalescence of multiple bubbles. Fig. 7 shows how individual, ‘naked’
DSPC bubbles coalesce into clusters of bubbles. This is easily
avoided by inclusion of PEGylated lipid (compare Fig. 7 with
Fig. 3), in which steric hindrance of polymer chains inhibits close
approach of adjacent bubbles. We find that 5 mol% PEG6000MS
gives very good, if not optimal, results, and have not yet investigated details of the PEG layer (such as brush layer versus mushroom).
We
do
note,
however,
that
a
PEGylated
phosphatidylcholine with a PEG molecular weight of 2000 was
ineffective in forming stable bubbles (we are unsure of the reason
for this but suspect it relates to chain unsaturation in the phospholipid rather than a de minimus PEG molecular weight; PEG2000MS
gave stable bubbles, although the PEG molecular weight did influ-
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7
Fig. 6. Bubble crumpling and phospholipid shedding upon gas dissolution: top panel: white arrows indicate a bubble as it undergoes crumpling upon gas leakage (left) and
just after resuming a (mostly) spherical shape and smooth surface upon shedding excess phospholipid (right). Bottom panel: a schematic drawing depicting crumpling and
phospholipid shedding, based on a previous model describing ‘‘zippering,” put forth by Borden and Longo [56].
Fig. 7. Bubble clustering: shown are successive intra-capillary images taken just seconds apart, demonstrating how DSPC bubbles, lacking any PEG, coalesce into clusters.
Panel a shows one such cluster. Bubble clusters travel as single entities, and panel b shows a second cluster shortly after it has arrived within the focal plane. Likewise, one
sees a third cluster, shortly after its arrival, in panel c.
ence bubble susceptibility to cavitation as compared with
PEG6000MS). Although not shown here, bubble coalescence could
also be avoided by inclusion of negatively charged phospholipids;
electrostatic forces would cause bubbles to repel upon close
approach.
3.2. Cavitation detection
Several means of detecting cavitation have been put forth
[7,15–17,57]. Primary among these are detection of free radicals
(typically peroxides) by chemical [58] and spectroscopic [59]
methods, respectively. It is known that inertial cavitation gives rise
to sub-harmonics (owing to a prolonged expansion phase with delayed collapse) and broadband noise [60]. As other phenomena
(e.g., non-linear bubble oscillations, surface waves, and bubble
state bifurcations) are known to also produce sub-harmonics, a
cavitation detection methodology based on sub-harmonics is not
advisable [61]. Sub-harmonics are necessary, but not sufficient,
indicators of inertial cavitation. However, broadband noise is –
for all practical purposes – exclusively an inertial cavitation-induced phenomenon and deemed a necessary and sufficient indicator of inertial cavitation. As a result, so-called passive cavitation
detection systems have been implemented, involving measurement of acoustic spectra and identification of the cavitation threshold as that pressure at which one observes a sudden onset of
broadband noise.
We favor this latter approach and demonstrate its utility here.
Fig. 8 shows raw data obtained from a typical (first version) cavitation experiment used to generate acoustic spectrograms. The horizontal axis tracks results of individual bubbles as an experiment
proceeds in time; each interval in time may be thought of as a repetition of identical experimental conditions, each with a new test
bubble. The ordinate is frequency of scattered acoustic radiation,
measured at the detection transducer; the intensity of radiation
is given on a relative grey scale (white being highest and black
being lowest). Accordingly, there are sporadic cases where one observes vertical black stripes; this indicates that no bubble was
present.
In Fig. 8a one sees an acoustic spectrogram obtained at a relatively low transmission intensity (50 kPa), one for which inertial
cavitation did not occur. In this instance, one sees re-radiation of
the primary incident wave at 1.8 MHz, evidenced by a bright and
horizontal white stripe on the spectrogram. Similarly, one observes
horizontal, white stripes parallel to, and at integer multiples of, the
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Fig. 8. Acoustic spectrograms for cavitation detection: (a) acoustic spectrogram obtained at 50 kPa of transmitting transducer. The primary transducer frequency is
observed as a bright, horizontal white line at 1.8 MHz. Also evident are harmonics (faint, horizontal white stripes) at multiples of the primary frequency; these are indicative
of non-linear processes, including non-linear wave propagation in water and non-linear bubble oscillations (i.e., stable cavitation); (b) acoustic spectrogram obtained at a
power of 1 MPa of transmitting transducer. The same phenomena observed in spectrogram a (50 kPa) also appear here. However, one also observes a sub-harmonic
(horizontal white stripe) at one half the primary frequencies. This is an indication, but not proof, of inertial cavitation. One also observes vertical, white stripes, which
constitute broadband noise, and this is deemed irrefutable evidence of inertial cavitation.
0.90
0.5% PEG6000
0.80
1% PEG6000
2% PEG6000
0.70
3% PEG6000
6% PEG6000
0.60
% Destroye
primary frequency. These harmonics are indicative of non-linear
bubble oscillations, or stable cavitation, and – to a lesser extent non-linear propagation of the ultrasound field in water.
The features just mentioned are also prominent in Fig. 8b,
which shows an acoustic spectrogram obtained at the highest
transmission intensity tested (1 MPa), and one for which inertial
cavitation is believed to have occurred. First note the appearance of
a horizontal, white stripe – albeit faint – at 0.9 MHz, exactly half of
the primary frequency. This sub-harmonic is the first clue that
inertial cavitation is present. However, for reasons mentioned previously, a sub-harmonic is not proof of inertial cavitation. Next
note the appearance, sporadically throughout the spectrogram, of
vertical, white stripes. These constitute broadband noise, and, in
our opinion, are direct evidence of inertial cavitation. Quite literally, a new tune is sung after each bubble cavitates or ‘‘pops.”
In addition to the spectrograms such as those in Fig. 8, it is possible to operate the cavitation detection system (second version) so
as to determine the percentage of bubbles destroyed over a wide
range of acoustic pressures. One such bubble destruction curve is
shown in Fig. 9. Here a cavitation threshold is easily observed as
an upturn in the bubble destruction curve. The purpose of Fig. 9
is not only to demonstrate the cavitation detection methodology
as applied to externally added microbubbles but also to test sensitivity of cavitation threshold to shell properties. In particular, we
prepared bubbles with varying amounts of PEG and with different
PEG molecular weights (PEG6000MS and PEG2000MS) as identified in Fig. 9.
The amount and molecular weight of PEG clearly influence bubble susceptibility to inertial cavitation. In the case of PEG6000MS,
the percent of bubbles destroyed is sensitive the mole fraction of
PEG in the range 0.5–3 mol%, higher PEG fractions giving greater
extents of bubble destruction. Addition of greater amounts of
PEG6000MS above 3 mol% (up to 9 mol%, the approximate maximal fraction one can incorporate within the bubble shell without
forming PEG aggregates) did not appear to impact cavitation
behavior. The influence of PEG2000MS was not as significant as
with PEG6000MS in that varying the PEG2000MS fraction did not
produce appreciable changes in the bubble destruction curve.
However, one observation is noteworthy: Whereas PEG2000MS
led to greater bubble destruction than PEG6000MS at mole fractions of 0.5 mol% and 1 mol%, PEG2000MS led to less bubble
destruction than PEG6000MS at mole fractions of 2 mol% and
3 mol%.
0.50
0.40
9% PEG6000
0.5% PEG2000
1% PEG2000
2% PEG2000
3% PEG2000
0.30
0.20
0.10
0.00
0.00
0.20
0.40
0.60
0.80
1.00
1.20
Acoustic Pressure, MPa
Fig. 9. Influence of shell property on cavitation threshold: the percentage of
bubbles destroyed due to inertial cavitation is plotted as a function of acoustic
pressure for two PEG molecular weights (2000 and 6000, in both cases the PEG is
linked via a monostearate) and a range of PEG mole fractions. Both the cavitation
threshold, which appears as a discontinuity on the abscissa, and the fraction of
bubbles destroyed, which takes on a sigmoidal profile with pressure, are sensitive
to changes in PEG mole fraction. In the case of PEG6000, however, little change is
noted for PEG fractions exceeding 3 mol%. Generally speaking, PEG6000MS impacts
bubble propensity toward cavitation more so than PEG2000MS. At PEG mole
fractions of 0.5% and 1 mol%, PEG6000MS leads to less cavitation than does
PEG2000MS; the opposite is true at PEG fractions of 2 mol% and 3 mol%.
The changes in cavitation threshold and bubble destruction are
most likely attributable to changes in bubble stiffness and accompanying changes in bubble resonance frequency. To be sure, bubble
resonance also changes due a change in mass as the fraction of PEG
increases, but this is a relatively small effect compared to stiffness
(the mass effect becomes more important for smaller bubbles). The
point is that for bubbles with a resonance frequency less than the
primary driving transducer frequency, an increase in the resonance
frequency associated with stiffening of the shell – as occurs with
addition of PEG – makes the bubbles more susceptible to cavitation
at the primary frequency. Although not shown here, one could
imagine just the opposite for bubbles with a primary frequency
greater than the primary driving frequency; that is, addition of a
stiffening agent would make the bubbles even less prone to cavitate. Shell stiffness is sometimes described and quantified by a
so-called shell parameter, Sp, which is related to the Young’s mod-
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Fig. 10. New drug delivery vehicle combining a giant uni-lamellar vesicle with a
microbubble: a giant, uni-lamellar vesicle (GUV, the larger, darker structure)
10 lm in diameter and containing a 50 mM glucose solution within the aqueous
core, is bound to an SF6-core/DSPC + PEG6000MS-shell microbubble (the smaller,
brighter object at 3 o’clock on the GUV). Binding was accomplished by incorporating biotinylated DSPE within both the GUV bilayer and the microbubble
monolayer and adding avidin to the aqueous medium. The aqueous medium
contained 50 mM sucrose to provide optical contrast of the GUV.
ulus [18,37,62,63]. Others have cast significant doubt on this approach [64,65], which is why we prefer casting the influence of
PEG in terms of rigorous colloidal science parameters describing
membranes and monolayers [19–25].
Despite current ambiguity over how best to model shell properties, a practical consequence of the correlation between cavitation
threshold and bubble resonance is the following: It is clear that the
extent of bubble cavitation can be tuned by tailoring bubble shell
chemistry. This suggests that a drug delivery vehicle could be
developed whose release rate is controllable with ultrasound. We
present such a vehicle in Fig. 10. The vehicle consists of a giant,
uni-lamellar vesicle, bound to a microbubble comprising an SF6
core and shell of DSPC and PEG6000MS. Binding was achieved
via biotinylated lipid in the bilayer and monolayer of the GUV
and microbubble, respectively, and avidin in the aqueous medium.
In a medical application, the GUV would contain a (either hydrophobic or hydrophilic) drug, which would be innocuous while contained within the GUV. Drug delivery would be controlled using
ultrasound as a remote, external trigger, in which cavitation (be
it stable or inertial) induces sonoporation of the GUV bilayer. As
cavitation, and therefore sonoporation, correlates with bubble
stiffness, the rate and extent of drug delivery can be controlled
by making appropriate changes in microbubble shell composition.
Acknowledgments
This research was supported in part by a Research Fellowship
from the Alexander von Humboldt Foundation.
SPW wishes to acknowledge and thank Mark Borden and Margie Longo for many useful discussions concerning microbubbles
and in particular for clarifying the issue of bubble dissolution in
saturated liquids mentioned in footnote 2.
References
[1] Postema M, Schmitz G. Bubble dynamics involved in ultrasonic imaging.
Expert Rev Mol Diagn 2006;6(3):493–502.
9
[2] Lum AFH, Borden MA, Dayton PA, Kruse DE, Simon SI, Ferrara KW. Ultrasound
radiation forces enables targeted deposition of model drug carriers loaded on
microbubbles. J Control Release 2006;111(1-2):128–34.
[3] Klibanov A. Targeted delivery of gas-filled microspheres, contrast agents for
ultrasound imaging. Adv Drug Deliv Rev 1994;37:139–57.
[4] Kheirolomoom A, Dayton PA, Lum AFH, Little E, Paoli EE, Zheng HR, et al.
Acoustically-active microbubbles conjugated to liposomes. Characterization of
a proposed drug delivery vehicle. J Control Release 2007;118(3):275–84.
[5] Unger EC, Matsunaga TO, McCreery T, Schumann P, Sweitzer R, Quigley R.
Therapeutic applications of microbubbles. Eur J Radiol 2002;42:160–8.
[6] Pong M, Umchid S, Guarino AJ, Lewin PA, Litniewski J, Nowicki A, et al.
Ultrasonics 2006;45:133–45.
[7] Leighton TG. The acoustic bubble. London: Academic Press; 1994.
[8] Marmottant P, Hilgenfeldt S. Controlled vesicle deformation and lysis by single
oscillating bubbles. Nature 2003;423:153–6.
[9] van Wamel A, Kooiman K, Harteveld M, Emmer M, Folkert JC, Versluis M, et al.
Vibrating microbubbles poking individual cells: Drug transfer into cells via
sonoporation. J Control Release 2006;112(2):149–55.
[10] Wu J, Ross JP, Chiu JF. Reparable sonoporation generated by microstreaming. J
Acoust Soc Am 2002;111(3):1460–4.
[11] Wu J. Theoretical Study on shear stress generated by microstreaming
surrounding contrast agents attached to living cells. Ultrasound Med Biol
2002;28(1):125–9.
[12] Schlicher RK, Radhakrishna H, Tolentino TP, Apkarian RP, Zarnitsyn V,
Prausnitz MR. Mechanism of intracellular delivery by acoustic cavitation.
Ultrasound Med Biol 2006;32(6):915–24.
[13] Greenleaf WJ, Bolander ME, Sarkar G, Goldring MB, Greenleaf JF. Artificial
cavitation nuclei significantly enhance acoustically induced cell transfection.
Ultrasound Med Biol 1998;24(4):587–95.
[14] Sundaram J, Mellein BR, Mitragotri S. An experimental and theoretical analysis
of ultrasound-induced permeabilization of cell membranes. Bioph J
2003;84:3087–101.
[15] Cramer E, Lauterborn W. Acoustic cavitation noise spectra. Appl Sci Res
1982;38:209–14.
[16] Hallow DM, Mahajan AD, McCutchen TE, Prausnitz MR. Measurement and
correlation of acoustic cavitation with cellular bioeffects. Ultrasound Med Biol
2006;32(7):1111–22.
[17] Ammi AY, Cleveland RO, Mamou J, Wang GI, Bridal SL, O’Brien Jr WD.
Ultrasonic contrast agent shell rupture detected by inertial cavitation and
rebound signals. IEEE Trans Ultrasonics Ferroelectrics Freq Control
2006;53(1):126–36.
[18] de Jong N, Hoff L, Skotland T, Bom N. Absorption and scatter of encapsulated
gas filled microspheres: theoretical considerations and some measurements.
Ultrasonics 1992;30(2):95–103.
[19] Evans DF, Wennerström H. The colloidal domain: where physics, chemistry,
biology, and technology meet. 2nd ed. New York: Wiley-VCH; 1999.
[20] Evans E, Rawicz W. Elasticity of ‘‘Fuzzy” biomembranes. Phys Rev Lett
1997;79(12):2379–82.
[21] Szleifer I, Gerasimov OV, Thompson DH. Spontaneous liposome formation
induced by grafted poly(ethylene oxide) layers: theoretical prediction and
experimental verification. Proc Natl Acad Sci USA 1998;95:1032–7.
[22] Rawicz W, Olbrich KC, McIntosh T, Needham D, Evans E. Effect of chain length
and unsaturation on elasticity of lipid bilayers. Biophys J 2000;79:328–39.
[23] Marsh D. Elastic constants of polymer-grafted lipid membranes. Biophys J
2001;81:2154–62.
[24] Rovira-Bru M, Thompson DH, Szleifer I. Size and structure of spontaneously
forming liposomes in lipid/PEG–lipid mixtures. Biophys J 2002;83:2419–39.
[25] Kim DH, Costello MJ, Duncan PB, Needham D. Mechanical properties and
microstructure of polycrystalline phospholipid monolayer shells: novel solid
microparticles. Langmuir 2003;19:8455–66.
[26] Apfel RE, Holland CK. Gauging the likelihood of cavitation from short-pulse,
low-duty cycle diagnostic ultrasound.
[27] Apfel RE. Acoustic cavitation prediction. J Acoust Soc Am 1981;69:1624–33.
[28] Flynn HG. Cavitation dynamics. II. Free pulsations and models for cavitation
bubbles. J Acoust Soc Am 1975;58:1160–70.
[29] Klibanov AL, Gu H, Wojdyla JK, Wible JH, Kim DH, Needham D, et al.
Attachment of ligands to gas-filled microbubbles via PEG spacer and lipid
residues anchored at the interface. In: Proc int symp control rel bioact mater,
vol. 26 (Revised July); 1999. p. 124–5.
[30] Chin CT, van Wamel A, Emmer M, de Jong N, Hall CS, Klibanov AL. Mechanisms
of ultrasonically-mediated drug delivery: high-speed camera observations of
microbubbles with attached ligands. In: Proc IEEE int ultrason symp; 2005.
[31] Mathivet L, Cribier S, Devaux PF. Shape change and physical properties of giant
phospholipid vesicles prepared in the presence of an AC electric field. Biophys J
1996;70:1112–21.
[32] Dimova R, Aranda S, Bezlyepkina N, Nikolov V, Riske KA, Lipowsky R. A
practical guide to giant vesicles. Probing the membrane nanoregime via optical
microscopy. J Phys Condens Matter 2006;18:S1151–76.
[33] Postema M, Mleczko M, Schmitz G. Experimental setup for synchronous
optical and acoustical observation of contrast microbubbles. Biomedizinische
Technik 2006;51(Suppl.):V75–76.
[34] Postema M, Mleczko M, Schmitz G. Contrast microbubble clustering at high
MI. In: Proc IEEE int ultrason symp; 2006. p. 1564–67.
[35] Mleczko M, Postema M, Schmitz G. Identifying nonlinear characteristics for the
bulk response of ultrasound contrast agents. In: Proc IEEE int ultrason symp;
2006. p. 1369–72.
Please cite this article in press as: Wrenn SP et al., Phospholipid-stabilized microbubbles: Influence of shell chemistry ..., Appl Acoust
(2008), doi:10.1016/j.apacoust.2008.09.017
ARTICLE IN PRESS
10
S.P. Wrenn et al. / Applied Acoustics xxx (2008) xxx–xxx
[36] Postema M, Schmitz G. Ultrasonic bubbles in medicine: influence of the shell.
Ultrasonics Sonochem 2007;14(4):438–44.
[37] Frinking PJA, de Jong N. Acoustic modeling of shell-encapsulated gas bubbles.
Ultrasound Med Biol 1998;24(4):523–33.
[38] Klibanov AL. Ligand-carrying gas-filled microbubbles: ultrasound contrast
agents for targeted molecular imaging. Bioconjugate Chem 2005;16:9–17.
[39] D’Arrigo JS. Physical Characteristics of ultra-stable lipid-coated microbubbles. J
Colloid Interfac Sci 1992;149(2):592–5.
[40] Klibanov AL, Rasche PT, Hughes MS, Wojdyla JK, Galen KP, Wible Jr JH,
Brandenburger GH. Detection of individual microbubbles of ultrasound
contrast agents: imaging of free-floating and targeted bubbles. Invest Radiol
2004;39:187–95.
[41] Zhao S, Borden M, Bloch S, Kruse D, Ferrara KW, Dayton PA. Radiation-force
assisted targeting facilitates ultrasonic molecular imaging. Mol Imaging
2004;3:135–48.
[42] Zhao Z, Liang HD, Mei XG, Halliwell M. Preparation, characterization, and
in vivo observation of phospholipid-based gas-filled microbubbles containing
Hirudin. Ultrasound Med Biol 2005;31(9):1237–43.
[43] Brown AC, Towles KB, Wrenn SP. Measuring raft size as a function of
membrane composition in PC-based systems: Part II – Ternary systems.
Langmuir 2007;23:11188–96.
[44] Brown AC, Towles KB, Wrenn SP. Measuring raft size as a function of
membrane composition in PC-based systems: Part I – Binary systems.
Langmuir 2007;23:11180–7.
[45] Troup GM, Wrenn SP, Apel-Paz M, Doncel GF, Vanderlick TK. A Time-resolved
diphenylhexatriene (DPH) anisotropy characterization of a series of model
lipid constructs for the sperm plasma membrane. Indus Eng Chem Res
2006;45(21):6939–45.
[46] Troup GM, Wrenn SP. Temperature and cholesterol composition-dependent
behavior of 1-myristoyl-2-[12-[(5-dimethylamino-1-naphthalenesulfonyl)amino]dodecanoyl]- sn -glycero-3-phosphocholine in 1,2-dimyristoyl- sn glycero-3-phosphocholine membranes. Chem Phys Lipids 2004;131:167–82.
[47] Wrenn SP, Gudheti MV, Veleva AN, Lee SP, Kaler EW. Characterization of model
bile using fluorescence energy transfer from dehydroergosterol to dansylated
lecithin. J Lipid Res 2001;42(6):923–34.
[48] Wrenn SP, Lee SP, Kaler EW. A fluorescence energy transfer study of lecithincholesterol vesicles in the presence of phospholipase C. J Lipid Res
1999;40(8):1483–94.
[49] Borden MA, Pu G, Runner GJ, Longo ML. Surface phase behavior and
microstructure of lipid/PEG-emulsifier monolayer-coated microbubbles.
Colloids and Surfaces B: Biointerfaces 2004;35:209–23.
[50] Kim DH, Costello MJ, Duncan PB, Needham D. Mechanical properties and
microstructure of polycrystalline phospholipid monolayer shells: novel solid
microparticles. Langmuir 2003;19:8455–66.
[51] Lin HY, Thomas JL. PEG-lipids and olig(ethylene glycol) surfactants enhance
the ultrasonic permeabilizability of liposomes. Langmuir 2003;19:1098–105.
[52] Simons K, Ikonen E. Functional rafts in cell membranes. Nature
1997;387:569–72.
[53] Jacobson K, Dietrich C. Looking at lipid rafts? Trends Cell Biol 1999;9:87–91.
[54] London E, Brown DA. Insolubility of lipids in Triton X-100: physical origin and
relationship to sphingolipid/cholesterol membrane domains (rafts). Biochim
Biophys Acta 2000;1508:182–95.
[55] Israelachvili J. Intermolecular and surface forces. 2nd ed. London: Academic
Press; 1992.
[56] Borden MA, Longo ML. Dissolution behavior of lipid monolayer-coated, airfilled microbubbles: effect of lipid hydrophobic chain length. Langmuir
2002;18:9225–33.
[57] Marin A, Sun H, Husseini GA, Pitt WG, Christensen DA, Rapoport NY. Drug
delivery in pluronic micelles: effect of high-frequency ultrasound on drug
release from micelles and intracellular uptake. J Control Release 2002;84:39–47.
[58] Kondo T, Misik V, Riesz P. Effect of gas-containing microspheres and echo
contrast agents on free radical formation by ultrasound. Free Rad biol med
1998;25:605–12.
[59] Alegria AE, Lion Y, Kondo T, Riesz P. Sonolysis of aqueous surfactant solutions –
probing the interfacial region of cavitation bubbles by spin trapping. J Phys
Chem 1989;93:4908–13.
[60] Neppiras EA. Acoustic cavitation. Physics Reports – Review Section of Physics
Letters 1980;61(3):159–251.
[61] Walton AJ, Reynolds GT. Sonoluminescence. Adv Phys 1984;33:595–660.
[62] de Jong N, Ten Cate FJ, Lancée CT, Roelandt JRTC, Bom N. Principles and recent
developments in ultrasound contrast agents. Ultrasonics 1991;29:324–30.
[63] de Jong N, Hoff L. Ultrasound scattering properties of Albunex microspheres.
Ultrasonics 1993;31(3):175–81.
[64] Doinikov AA, Dayton PA. Spatio-temporal dynamics of an encapsulated gas
bubble in an ultrasound field. J Acoust Coc Am 2006;120(2):661–9.
[65] Doinikov AA, Dayton PA. Maxwell rheological model for lipid-shelled ultrasound
microbubble contrast agents. J Acoust Soc Am 2007;121(6):3331–40.
[66] Epstein PS, Plessett MS. Gas bubbles in solutions. J Chem Phys
1950;18(11):1505–9.
[67] Duncan PB, Needham D. Test of the Epstein–Plesset Model for gas
microparticle dissolution in aqueous media: effect of surface tension and gas
undersaturation in solution. Langmuir 2004;20:2567–78.
Please cite this article in press as: Wrenn SP et al., Phospholipid-stabilized microbubbles: Influence of shell chemistry ..., Appl Acoust
(2008), doi:10.1016/j.apacoust.2008.09.017