# Antibubbles enable tunable payload release with low-intensity ultrasound

###### Abstract

The benefits of ultrasound are its ease-of-use and its ability to precisely
deliver energy in opaque and complex media. However, most materials responsive
to ultrasound show a weak response, requiring the use of high powers, which
are associated with undesirable streaming, cavitation, or temperature rise.
These effects hinder response control and may even cause damage to the medium
where the ultrasound is applied. Moreover, materials that are currently in use
rely on all-or-nothing effects, limiting the ability to fine-tune the response
of the material on the fly. For these reasons, there is a need for materials
that can respond to low intensity ultrasound with programmable responses. Here
it is demonstrated that antibubbles are a low-intensity-ultrasound-responsive
material system that can controllably release a payload using acoustic
pressures in the kPa range. Varying their size and composition tunes the
release pressure, and the response can be switched between a single release
and stepwise release across multiple ultrasound pulses. Observations using
confocal and high-speed microscopy revealed different ways that can lead to
release. These findings lay the groundwork to design antibubbles that
controllably respond to low-intensity ultrasound, opening a wide range of
applications ranging from ultrasound-responsive material systems to carriers
for targeted delivery.

###### keywords:

antibubble, smart-material, payload, triggered release, low-intensity
ultrasound

Nicolas Moreno-Gomez*† Athanasios G. Athanassiadis*† Albert T. Poortinga Peer
Fischer

† These authors contributed equally.

Dr. Nicolas Moreno-Gomez
Address: Institute for Molecular Systems Engineering and Advanced Materials,
Im Neuenheimer Feld 225, 69120 Heidelberg, Germany and Max Planck Institute
for Medical Research, Jahnstr. 29, 69120 Heidelberg, Germany.
Email Address<EMAIL_ADDRESS>

Dr. Athanasios G. Athanassiadis
Address: Institute for Molecular Systems Engineering and Advanced Materials,
Im Neuenheimer Feld 225, 69120 Heidelberg, Germany and Max Planck Institute
for Medical Research, Jahnstr. 29, 69120 Heidelberg, Germany.
Email Address<EMAIL_ADDRESS>

Dr. Albert T. Poortinga
Address: Polymer Technology group, Eindhoven University of Technology,
Eindhoven, The Netherlands
De Rondom 70, 5612 AP Eindhoven, The Netherlands

Prof. Dr. Peer Fischer
Address: Institute for Molecular Systems Engineering and Advanced Materials,
Im Neuenheimer Feld 225, 69120 Heidelberg, Germany and Max Planck Institute
for Medical Research, Jahnstr. 29, 69120 Heidelberg, Germany.

##  1 Introduction

Ultrasound is a versatile tool in diverse fields, ranging from chemistry and
fabrication[1, 2, 3], to biology and medicine [4]. Recent advances in
ultrasound-responsive materials as well as advances in physical acoustics have
expanded the settings in which ultrasound can be used and the capabilities
that it can provide [5, 6]. In particular, the ability to precisely shape
ultrasound fields [7] has made it possible to remotely trigger responses in
localized regions on demand [2], while benefiting from ultrasound’s unique
characteristics compared to other methods of remote stimulation. For instance,
ultrasound provides higher spatial resolution than magnetics, while
propagating further than light through complex and opaque media [8, 9, 10].
Additionally, ultrasound can drive different physical phenomena – via acoustic
radiation forces, cavitation or streaming – that can be exploited when
developing new responsive systems [5]. Nevertheless, despite the benefits of
ultrasound, there are few materials that can be triggered or controlled with
ultrasound.

Most ultrasound-responsive materials require moderate- or high-intensity
ultrasound to trigger a desired effect [5, 11]. Examples of this mostly arise
in controlled drug delivery, where the carrier, often a microbubble, is
stimulated using ultrasound to release a drug and initiate a therapeutic
effect [12]. Beyond drug delivery there are examples where ultrasound-
responsive systems are used to induce self-healing in construction materials
[13, 14] or for in-vivo fabrication of hydrogels through external ultrasound
stimulation [15], demonstrating the importance of controlling the payload
release for different applications. In many examples, high-intensity
ultrasound is required to release a payload from a carrier such as
microbubbles [16], liposomes [17, 18], or phase-change droplets [19] using
acoustic pressures on the order of MPa and frequencies of tens of kHz up to a
few MHz [5, 12, 20, 21, 22, 23].

However, the use of high-intensity ultrasound can lead to undesirable effects
that limit its application in sensitive material or biological systems.
Thermal, mechanical, and chemical damage is possible within the ultrasound
beam through absorption, cavitation, and sonochemical effects. The onset of
such effects are dependent on a combination of intensity (pressure),
frequency, and exposure time. Commonly adopted thresholds for high-intensity
ultrasound correspond to pressures of $150\text{\,}\mathrm{kPa}$ (time-
average) and $2\text{\,}\mathrm{MPa}$ (peak) at $1\text{\,}\mathrm{MHz}$ in
water [24]. However, detrimental effects in materials have been observed at
pressures as low as $170\text{\,}\mathrm{kPa}$ at $1\text{\,}\mathrm{MHz}$
[25], and certain systems can be much more susceptible to damage than others
(e.g. in ophthalmology [24, 26]). In order to be useful in a wide range of
contexts without inducing unwanted damage, there is therefore a need for
ultrasound-responsive materials that respond to low-intensity ultrasound,
ideally in the range of $200\text{\,}\mathrm{kPa}$ or below.

An additional challenge when using existing ultrasound-responsive systems is
their release response. There is typically a narrow range of excitation
pressures, above which the carrier is destroyed and the payload is released in
a single event [9, 16, 27]. While such behavior might be desirable in some
applications, e.g. for rapid delivery of a payload, there are other settings
where more control is required. For instance, when the payload should be
precisely dosed in response to real-time feedback [11], or when the payload
needs to be slowly delivered over an extended period of time [28]. In such
settings, existing ultrasound-responsive carriers do not provide adequate
solutions.

In this work, we identify Pickering-stabilized antibubbles as a new carrier
for triggered release and show that they respond robustly to low-intensity
ultrasound. Antibubbles consist of one [29, 30, 31] or more liquid droplets
[32, 33] surrounded by a gas layer. This structure is not naturally
thermodynamically stable [34], but can be stabilized using fumed silica
particles (Pickering stabilization) instead of surfactant molecules [29, 32,
35]. Unlike microbubble-based carriers, which require specially-modified
payloads that can be attached to the outer shell [36], the internal droplets
in an antibubble can carry large volumes of payload without special
preparation. Because of these benefits, antibubbles are a new class of
ultrasound contrast agents that have recently been proposed for ultrasonic
drug delivery [33].

Here we demonstrate, for the first time, that payload delivery via antibubbles
can be triggered by low-intensity ultrasound with controllable temporal
release profiles. Moreover, we show that the release response can be shifted
to lower pressures, and can exhibit different qualitative behaviors by tuning
the formulation. We use optical fluorescence measurements to detect the
release of the payload as a function of ultrasound exposure and antibubble
composition. We find that, unlike with existing carriers, it is possible to
change the release behavior from single release to multiple-release by varying
the composition of the antibubble. Similarly, the release pressure can be
adjusted via composition variation. Finally, we use high-speed brightfield
microscopy to explore the origins of the different release mechanisms. Our
results open the door to designing tailored antibubble formulations for
specific applications, making antibubbles a valuable and versatile component
for the design of new low-intensity ultrasound-activated smart materials.

##  2 Results and Discussion

###  2.1 Antibubble Fabrication and Characterization

Antibubbles are fabricated [33] by creating a water-in-oil-in-water (W/O/W)
Pickering emulsion template with the desired payload dissolved in the inner
water phase. The solvent is then extracted by freeze-drying, creating a stable
powder template with a very long shelf life. Before use, the antibubbles are
reconstituted from the powder in an aqueous solution of sodium chloride. The
fabrication process is schematically illustrated in Figure 1a along with a
confocal image of a single antibubble to illustrate their structure (also see
video S1). For this image, we incorporate a fluorescent calcein dye as a
payload at concentration of 2.6 mM in order to visualize the core structure.
These images confirm that multiple droplets are encapsulated in the core, and
that these tend to sediment toward the bottom of the antibubble because of
their higher density compared to the gas phase [30].

As has been previously described for other carriers [17], to study and
quantify the release from antibubbles with ultrasound, we increase the
concentration of the fluorescent calcein to 119 mM to use the self-quenching
of calcein at high concentrations (Figure S1) to distinguish between intact
antibubbles which are characterized by low fluorescence levels when the dye is
encapsulated, and antibubbles from which a payload is released. After release,
the calcein is diluted in the surrounding fluid, quenching ceases, and
fluorescence is observed.

The calcein-loaded inner phase of the antibubble template is prepared in an
aqueous solution and also includes sodium hydroxide to enhance the calcein
solubility, as well as maltodextrin as a stabilizer for freeze-drying. Small
droplets of concentrated calcein are then formed by combining the aqueous
solution with a dispersion of hydrophobic fumed silica in cyclooctane. This
water-in-oil (W/O) emulsion is formed by vigorously stirring the two
components together using an ultrasonic homogenizer (unrelated to the
ultrasonic transducers used in the experiments below). The W/O emulsion is
further homogenized with an aqueous dispersion of fumed silica, maltodextrin,
mannitol and sodium chloride to produce the final water-in-oil-in-water
(W/O/W) template. The additives are included to balance the osmotic pressure,
thereby preventing the inner droplets from swelling or deswelling. The W/O/W
template is formed by rotor-stator homogenization. It is important to note
that the final size of the antibubbles is largely dictated by the
homogenization rate used in this step. Next, the template is washed and
concentrated using centrifugation: the positively-buoyant W/O/W droplets rise
to the top without coalescing, and can be separated from any free
(unencapsulated) calcein. The concentrates obtained after centrifugation are
freeze-dried to replace the intermediate oil phase with air. During this step,
it is important to avoid structure collapse that can occur if the sample melts
before starting the drying process. Therefore, we freeze the sample at
temperatures below the glass transition temperature, which in our case is
controlled by the concentration of maltodextrin in the internal and external
aqueous phases. The concentrates were frozen in a round bottom glass flasks
for 6 h in a freezer at -60∘C, a temperature low enough to prevent melting of
the structure before starting the drying process. The samples were then dried
by connecting the round-bottom flasks to a 0.1 mbar vacuum line for 24 h.
During this time, the cyclooctane and most of the water are removed to produce
a dried powder that can be stored for long periods of time. Finally,
antibubbles are formed by reconstitution of the powder in an aqueous solution
of sodium chloride.

Figure 1: Fabrication and characterization of antibubbles. (a) Antibubbles
were fabricated by freeze-drying a double emulsion template with multiple
cores containing calcein, as visualized using Confocal microscopy (scale bar
corresponds to $15.0\text{\,}\mathrm{\SIUnitSymbolMicro m}$. (b)-(e) Both
homogenization rate and fumed silica composition play a role in determining
the size of the antibubbles. Hydrophilically-coated antibubbles obtained after
homogenization (Rotor-stator) at (b) 7000 rpm and (c) 14000 rpm show a small
decrease in size with shear rate, while hydrophobically-coated antibubbles
obtained after homogenization at (d) 7000 rpm and (e) 14000 rpm vary in size
more significantly. (f) In fluorescence images the reconstituted antibubbles
(here: hydrophilic, $51\text{\,}\mathrm{\SIUnitSymbolMicro m}$) can be
identified by their dark cores surrounded by a gas layer. The dark cores
reflect the quenched calcein, while free calcein in the background solution
provides the ambient green fluorescence.

We fabricated and characterized different antibubble formulations to test
their response to ultrasound. In analogy to microbubbles [37], we hypothesized
that size and interfacial energy dictate the acoustic response of antibubbles.
Therefore, four different formulations were prepared by varying the size of
the template and the composition of the outer shell, while keeping the inner
cores the same for all formulations. The sizes are varied by homogenizing the
W/O/W emulsion at either 7000 or 14000 rpm. The outer shell is varied by using
two distinct types of commercially-available fumed silica particles. One type
(Aerosil 200) was predominantly hydrophilic, while the other (Aerosil R972)
has a modified surface with carbon content of 0.6-1.2$\%$ [38], making it
predominantly hydrophobic. The first formulation we use in experiments
consists of the hydrophilic Aerosil 200 in a 2:1 mass ratio with Aerosil R972,
and is thus referred to as ‘hydrophilic’ below. The second formulation
consists of only hydrophobic Aerosil R972 fumed silica and is referred to as
‘hydrophobic’ below. As shown in Figures 1b-e, both formulations produce a
similar size distribution when homogenized at 7000 rpm. However, at 14000 rpm
the hydrophobic formulation produces much smaller antibubbles. All
formulations show good stability over time (see SI Figure S2) by retaining
their size for a period of 20 h. Quenching of inner fluorescence is indicated
by the appearance of the dark antibubble centers in both brightfield and
fluorescence microscopy (Figure 1f and supporting video S2). Free calcein in
the external aqueous phase is removed via further washing of the antibubbles:
the buoyant antibubbles are collected from the top of the aqueous solution and
transferred to a new sodium chloride solution during the washing steps.

###  2.2 Release Triggered by Low Intensity Ultrasound

To determine the response of antibubbles to ultrasound, we measured the change
in fluorescence caused by the release of calcein after exposure to ultrasound.
The experimental setup is shown in Figure 2a. Antibubbles were observed in a
quartz spectroscopy cuvette that had an ultrasonic transducer bonded to one
side of the cuvette. Fluorescence was measured by illuminating the sample with
460 nm light from a fiber-coupled LED, and collecting the emitted light with a
fiber-coupled spectrometer oriented 90∘ to the excitation fiber. The
fluorescence intensity at 520 nm, corresponding to the calcein emission peak,
was used as an indicator of antibubble bursting.

Because the antibubbles are positively buoyant, the sample was mixed by gently
shaking the cuvette before each fluorescence measurement and ultrasound
exposure. Separate measurements confirmed that the antibubbles do not burst
during gentle shaking. Before any ultrasound exposure, a baseline fluorescence
level was measured. Then, the sample was gently shaken and exposed to an
ultrasound pulse at 1 MHz for 2 s. This process was repeated with increasing
ultrasound pressures to identify when the antibubbles release the payload.
Fluorescence signals were collected continuously during and for 1 min after
each ultrasound exposure to confirm the signal stability. The ultrasound
response was measured using at least three different samples of each
antibubble formulation. Because absolute fluorescence intensities vary from
sample to sample, the resulting signal was normalized after background
subtraction (see SI Figure S3 for raw fluorescence data). The normalized
fluorescence intensity is plotted as a function of acoustic pressure for each
formulation in Figures 2b-e. Our measurements reveal that the antibubble
response to ultrasound is strongly dependent on the outer shell composition
and size. The results are plotted in Figure 2 and summarized in Table 1. This
opens up the possibility to tune the release characteristics in these
carriers.

Figure 2: Experimental setup and results for fluorescence measurements as a
function of ultrasound exposure. (a) Antibubbles were loaded into a cuvette
and exposed to ultrasound using an integrated transducer at 1 MHz.
Fluorescence signals at 520 nm were measured using a fiber-coupled
spectrometer, oriented perpendicular to the excitation light (460 nm). (b)-(e)
The antibubble response to ultrasonic pressure depends on antibubble
composition. The pressure response for (b)
$51\text{\,}\mathrm{\SIUnitSymbolMicro m}$ and (c)
$34\text{\,}\mathrm{\SIUnitSymbolMicro m}$ hydrophilic antibubbles both
demonstrate a clear threshold behavior with fluorescence intensities
saturating above the threshold. The response of (d)
$58\text{\,}\mathrm{\SIUnitSymbolMicro m}$ and (e)
$19\text{\,}\mathrm{\SIUnitSymbolMicro m}$ hydrophobic antibubbles show
differing behaviors, with a low-intensity threshold only observable in the
smaller antibubbles. Markers represent measurements on independent samples,
the solid line represents the averaged intensity and the shading indicates one
standard deviation. (f) The hydrophilic antibubbles release all the
fluorescence after a single ultrasound pulse, with no significant release
after subsequent pulses at the same pressure (140 kPa). (g) The hydrophobic
antibubbles release the core fluorescence incrementally across multiple pulses
at the same pressure (70 kPa).

Antibubbles with hydrophilic outer shells demonstrate single-release behavior,
with the release pressure dependent on the size. The larger
($51\text{\,}\mathrm{\SIUnitSymbolMicro m}$) hydrophilic antibubbles (Figure
2b) release calcein already at a low pressure of $88\text{\,}\mathrm{kPa}$
(equivalent intensity $258\text{\,}\mathrm{mW}\text{\,}{\mathrm{cm}}^{-2}$),
with fluorescence release increasing to saturation at
$109\text{\,}\mathrm{kPa}$. The smaller
($34.0\text{\,}\mathrm{\SIUnitSymbolMicro m}$) antibubbles (Figure 2c)
demonstrated a similar release behavior, yet starting from a lower pressure
below $7\text{\,}\mathrm{kPa}$ (equivalent intensity
$2\text{\,}\mathrm{mW}\text{\,}{\mathrm{cm}}^{-2}$) , with the amount of
calcein release increasing until $53\text{\,}\mathrm{kPa}$, where the
fluorescence signal saturates, indicating no further release from the core.
These differences in the response indicate that by changing the size, the
required acoustic pressure for release can be shifted.

To confirm that the observed fluorescence release was due solely to the
pressure level and not to the repeated exposures of ultrasound, we repeated
our measurements for select pressures with new samples for each pressure
(indicated by cross markers in Figure 2b; 5 independent samples). The amount
of fluorescence measured in these independent experiments matches closely with
the previously measured values. As a final demonstration that multiple
exposures do not significantly affect the measurements for hydrophilic
antibubbles, we exposed a single sample of the large hydrophilic antibubbles
to multiple pulses at $140\text{\,}\mathrm{kPa}$, which is above their
bursting pressure. As shown in Figure 2f, the fluorescence increases
drastically after the first ultrasound pulse, with minimal further release of
calcein after subsequent pulses. These measurements confirm that pressure
plays the dominant role in determining release for the hydrophilic
antibubbles, and not multiple exposures. These results also indicate that the
antibubbles are stable against ultrasound exposure at pressures below the
bursting pressure.

In contrast to the hydrophilic antibubbles, those with the hydrophobic shell
demonstrate more varied size-dependent behavior in response to ultrasound. As
shown in Figure 2d, the larger ($58\text{\,}\mathrm{\SIUnitSymbolMicro m}$)
antibubbles do not release appreciable amounts of calcein after irradiation.
The same conclusion can be drawn from the raw fluorescence signal (Figure
S3c). The formulation with a smaller diameter
($19.0\text{\,}\mathrm{\SIUnitSymbolMicro m}$), however, shows that above
$49\text{\,}\mathrm{kPa}$ (equivalent intensity
$80\text{\,}\mathrm{mW}\text{\,}{\mathrm{cm}}^{-2}$) the fluorescence
intensity increases linearly with each ultrasound pulse to at least
$250\text{\,}\mathrm{kPa}$ (Figure 2e), indicating an ongoing release process
that does not saturate. To test this behavior further, we measured the calcein
released by these small, hydrophobic antibubbles across multiple pulses at
$70\text{\,}\mathrm{kPa}$ (Figure 2g). We find that even at a single
excitation pressure, the antibubbles release small amounts of calcein after
each pulse, reflecting a stepwise release process rather than a single one.

Table 1: Response of different antibubble formulations to ultrasonic excitation at 1 MHz.  Shell Composition† | Homogenization Rate | Size | Release Type | Release Threshold
---|---|---|---|---
Hydrophilic | $14\,000\text{\,}\mathrm{r}\mathrm{p}\mathrm{m}$ | $34\,[8]\text{\,}\mathrm{\SIUnitSymbolMicro m}$ | Single | $<7\text{\,}\mathrm{kPa}$
Hydrophilic | $7000\text{\,}\mathrm{r}\mathrm{p}\mathrm{m}$ | $51\,[13]\text{\,}\mathrm{\SIUnitSymbolMicro m}$ | Single | $88\,[7]\text{\,}\mathrm{kPa}$
Hydrophobic | $14\,000\text{\,}\mathrm{r}\mathrm{p}\mathrm{m}$ | $19\,[5]\text{\,}\mathrm{\SIUnitSymbolMicro m}$ | Multi | $49\,[14]\text{\,}\mathrm{kPa}$
Hydrophobic | $7000\text{\,}\mathrm{r}\mathrm{p}\mathrm{m}$ | $58\,[17]\text{\,}\mathrm{\SIUnitSymbolMicro m}$ | None | $>250\text{\,}\mathrm{kPa}$
$\dagger$ Hydrophobic - Aerosil R927; Hydrophilic - Aerosil 200 and Aerosil
R972 (2:1)

In addition to the fluorescence measurements, we checked how ultrasound
exposure alters the size distribution of antibubbles. We sampled hydrophilic
antibubbles from a single batch before and after exposure to
$140\text{\,}\mathrm{kPa}$ ultrasound and measured their sizes using
brightfield microscopy and standard image processing tools (SI Figure S4).
These measurements show that not all antibubbles burst. Rather, the mean
diameter of the antibubbles shifts from $51\text{\,}\mathrm{\SIUnitSymbolMicro
m}$ to $45\text{\,}\mathrm{\SIUnitSymbolMicro m}$ ($P<0.05$, $n=300$). In
addition, the fraction of antibubbles larger than
$50\text{\,}\mathrm{\SIUnitSymbolMicro m}$ decreased while the fraction of
smaller antibubbles increased. This observation suggests that antibubbles with
a diameter larger than $50\text{\,}\mathrm{\SIUnitSymbolMicro m}$ are more
sensitive to the $1\text{\,}\mathrm{MHz}$ ultrasound. However, given the
observations of multiple release in the smaller hydrophilic antibubbles it is
still unclear whether the smaller hydrophobic antibubbles can release calcein
without being completely destroyed. Similarly, it is not clear whether the
change in size distribution is due to complete destruction of larger
antibubbles, or fragmentation that produces smaller bubbles.

Our results clearly demonstrate that the response of antibubbles to ultrasound
is strongly dependent on outer shell composition and antibubble size. While
the hydrophilic antibubbles release the core contents in a single bursting
event upon ultrasound exposure, the hydrophobic antibubbles release less of
their core contents per exposure, providing on-demand dosing of the payload.
In both cases, the bubble sizes determine the pressure threshold, above which
the antibubbles begin releasing their contents. This behavior, however also
shows different trends depending on the shell composition. For the hydrophilic
antibubbles, smaller bubbles exhibit a higher release threshold, while for the
hydrophobic antibubbles, the larger bubbles do not exhibit any release
behavior within the range of pressures investigated. These opposing trends
raise interesting questions for the role of size in antibubble release. By
analogy with microbubbles, one might expect resonant effects to play an
important role, with bubbles resonant closer to the excitation frequency
requiring lower pressures for release than bubbles further from resonance.
However by this logic the smaller antibubbles should have released at lower
pressures compared with the larger ones when excited at
$1\text{\,}\mathrm{MHz}$ [39], which is not the case for the hydrophilic
antibubbles. These results indicate a more complex interplay between size and
shell composition in determining the acoustic response of antibubbles.

The multi-release behavior and low pressure thresholds make antibubbles more
versatile tools than existing carriers. Most existing carriers release their
payload all at once [5], with only a few capable of gradual release [11, 40,
41, 42, 43, 44]. One of the few examples where two different release
mechanisms have been observed is in polymeric micelles. Depending on the type
of polymer used and the ultrasound parameters (frequency, amplitude), single
release was possible after irreversible disruption of the micelle structure,
or partial release could be caused by temporal disruption of the self-
assembled structure [41]. Since each of these responses is caused by different
phenomena, it is necessary to use different ultrasound amplitudes for each
polymer structure. By contrast, antibubbles exhibit the two different
behaviors under similar ultrasound conditions and only require different
composition of the external shell. This makes antibubbles one of the few
carriers where the release can be tuned. Interestingly, the release profile
can be adjusted using the same ultrasound settings. Furthermore, much lower
acoustic pressures are required to release the contents of the antibubbles
compared to other carriers like liposomes (1.5-2.0 MPa)([18]), microbubbles
(0.5-5.0 MPa) [16] or PFC-droplets (0.3-8.5 MPa) [19]. The release pressures
for antibubbles
($7\text{\,}\mathrm{kPa}49\text{\,}\mathrm{kPa}88\text{\,}\mathrm{kPa}$)
correspond to low intensities
($2\text{\,}\mathrm{mW}\text{\,}{\mathrm{cm}}^{-2}80\text{\,}\mathrm{mW}\text{\,}{\mathrm{cm}}^{-2}258\text{\,}\mathrm{mW}\text{\,}{\mathrm{cm}}^{-2}$)
and low mechanical indices (0.01, 0.05, and 0.09 for $1\text{\,}\mathrm{MHz}$
excitation). These low pressures preclude nonlinear propagation in the
background medium and ensure that destructive thermal or cavitation effects do
not pose a risk to surrounding media. The low release threshold is also
specific to ultrasound, and the bubbles remain stable even in the presence of
mild mechanical agitation such as flows and stirring. While higher background
ultrasound intensities could spuriously trigger unspecific release, targeted
ultrasonic fields typically localize the ultrasonic pressure sufficiently and
therefore should not trigger release beyond the higher-intensity focus.
Between the specificity of the the release to ultrasonic stimulation and the
low release thresholds, antibubbles can be a very attractive candidate for
applications requiring tunable, on-demand payload release in sensitive
environments.

###  2.3 Antibubble bursting dynamics and release mechanisms

The ability to tune antibubble properties for use in different settings
requires a better understanding of the release mechanism for individual
antibubbles, and how this mechanism varies with size and composition. As
discussed above, resonant effects are expected to play a role, however this is
likely in a complex interplay with other effects stemming from the surface
energy of the antibubbles. Other effects, such as direct stretching of the
fluid and antibubbles in the acoustic waves, can be ruled out as the
$\sim$100\text{\,}\mathrm{kPa}$$ pressures used above produce a peak molecular
displacement of $11\text{\,}\mathrm{nm}$. Taken over the acoustic wavelength
($1.5\text{\,}\mathrm{mm}$ at $1\text{\,}\mathrm{MHz}$), this corresponds to a
peak strain on the order of $0.0007\text{\,}\mathrm{\char 37\relax}$ \- far
too small to have a direct effect on antibubble release.

In order to clarify what happens at a single-bubble level during ultrasound
exposure, we made use two different microscopy techniques. First, we measured
the fluorescence released from individual antibubbles using confocal
microscopy. Second, to observe the dynamics of antibubble release, we recorded
the response of antibubbles to ultrasound using high-speed brightfield
microscopy. These two imaging techniques provide us complementary information
on the single-antibubble release dynamics: confocal imaging allows us to
assess release and bursting after an entire pulse, while the high-speed
imaging reveals antibubble dynamics on short timescales associated with each
ultrasound oscillation. In these experiments we only investigated the large
hydrophilic antibubbles, since the better-defined pressure threshold allows
for clearer insights into the release process.

For the microscopy measurements, we used a glass-plate excitation setup, which
allows for transmission imaging of the sample in a small chamber (Figure 3a).
The antibubble sample was suspended in a solution of 3. wt$\%$ of NaCl 0.3
wt$\%$ xantham gum to minimize antibubble motion and aggregation during the
excitation. A low-concentration sample was loaded into a well (Gene Frame,
Thermo Scientific) on a glass plate, and the well was sealed with a
transparency film. The sample was excited by a piezoelectric transducer glued
to the glass plate, which was driven by $300\text{\,}\mathrm{ms}$ pulses at
the radial resonance frequency of the transducer
($90.5\text{\,}\mathrm{kHz}$). This excitation mode transmitted acoustic
pulses through the glass into the sample chamber (SI Figure S5). The lower
frequency was used to permit for direct observation of the dynamics via high-
speed imaging.

Figure 3: Observation of payload release from individual hydrophilic
antibubbles using confocal fluorescence microscopy. Panel (a) illustrates the
glass-slide setup used for the observation, where antibubbles suspended in a
medium of xanthan gum are placed inside a chamber. Panel (b) shows the
antibubbles before and after ultrasound stimulation. The release of the
payload is confirmed by the detection of increased fluorescence locally where
the antibubbles were located. Signs of antibubble fragmentation and
destruction are visible as trails of silica, alongside bubbles that have
released fluorescence but otherwise remained intact.

The confocal measurements revealed clear release from individual antibubbles,
as well as signs of fragmentation and partial release. As shown in Figure 3b,
intact antibubbles before ultrasound exposure can be identified by their
compact fluorescent cores. After a single 300 ms ultrasound pulse, calcein was
visibly released and slowly diffused through the surrounding medium. A closer
look at individual antibubbles shows evidence of motion in the ultrasound
field and large fragments (most likely silica remaining after antibubble
fragmentation) still emitting fluorescence, as well as some intact antibubbles
with a weakly fluorescent core. This suggests that the antibubbles respond
differently to ultrasound. While some antibubbles were completely destroyed
during the pulse, others moved under the influence of radiation forces,
leaving a trail of the fumed silica and releasing at least part of their
payload. Thus intact bubbles or antibubbles after exposure also contribute to
the observed fluorescence change, suggesting that the payload release does not
require complete destruction of the hydrophilic antibubbles, but can also
involve more complicated dynamics such as fragmentation or transient bridging
of the core with the external fluid.

Figure 4: Time-resolved response of four antibubbles to ultrasound recorded
with high-speed microscopy. In panel (a), three antibubbles of varying sizes
exhibit distinct behaviors: antibubble A
($72\text{\,}\mathrm{\SIUnitSymbolMicro m}$) undergoes highly non-linear
oscillations, antibubble B ($47\text{\,}\mathrm{\SIUnitSymbolMicro m}$) sheds
material and interacts with an oscillating bubble cluster, and antibubble C
($34\text{\,}\mathrm{\SIUnitSymbolMicro m}$) ejects material during stable
oscillations. Panel (b) shows a fourth antibubble
($32\text{\,}\mathrm{\SIUnitSymbolMicro m}$) that is similar in size to
antibubble C, which releases material from the core after 15 ms, as indicated
by the sudden brightening of the core due to an increase in the visibility of
transmitted light through the bubble. The antibubble subsequently undergoes
much larger oscillations. Frames in panel (a) were extracted from supporting
Video S3 and panel (b) from Video S4. Scale bars represent
$50\text{\,}\mathrm{\SIUnitSymbolMicro m}$.

We directly observed multiple individual antibubbles using high-speed
microscopy and identified three characteristic responses that could lead to
payload release. The antibubble sample was again transferred to the glass
plate, placed in an upright microscope and recorded with 10$\times$
magnification at 120000 frames per second using a high-speed camera. The
characteristic responses are exemplified by three different antibubbles with
different sizes that were observed simultaneously during ultrasound exposure
(diameters: A - $72\text{\,}\mathrm{\SIUnitSymbolMicro m}$, B -
$47\text{\,}\mathrm{\SIUnitSymbolMicro m}$, and
C-$34\text{\,}\mathrm{\SIUnitSymbolMicro m}$). Figure 4a shows select frames
extracted from supporting video S3, illustrating the response of three
antibubbles. Antibubble A is the first to start oscillating, followed by
antibubbles B and C. The oscillations of antibubble A become larger with time
and quickly become nonlinear, exhibiting higher order oscillations. Antibubble
B, of intermediate size, oscillates until it begins to release material,
probably silica, followed by its destruction after interacting with a bubble
cluster, likely formed by other antibubbles out of the frame. Here, the
release of the payload is not only due to the collapse of the structure, but
also involves interactions with other antibubbles and with material coming
from the fragmentation of other structures. Lastly, antibubble C oscillates
weakly at the start of the ultrasound pulse. However, after more time
(supporting video S3), the oscillations of antibubble C become larger and it
releases its payload while remaining intact, revealing a third release
mechanism. The same mechanism was observed in a fourth antibubble
($32\text{\,}\mathrm{\SIUnitSymbolMicro m}$ diam.), shown in Figure 4b and
supporting video S4. For these antibubbles, linear ultrasound-induced
oscillations resulted in an eventual release of the payload, marked by a
transition from opaque dark cores to transparent ones. This process,
accompanied by a slow movement of the antibubble, continues with the ejection
of material (likely silica), and is followed by large oscillations that would
not be possible with an intact core. Finally, the residual microbubble jets
away, leaving a trace similar to what was observed in the confocal images
(Figure 3b).

Our microscopy results reveal that individual antibubbles can demonstrate a
rich variety of behaviors leading to payload release. Antibubbles can release
during a destructive collapse or fragmentation, which are processes likely
driven by large nonlinear oscillations or interactions with other bubbles or
antibubbles. The fact that nonlinear oscillations can onset so quickly in low
pressures is likely a result of two factors. Most importantly is the role of
the core. Since the core is incompressible and typically around 80% of the
antibubble diameter, there is a smaller volume of air that can participate in
oscillations, and interactions between the inner and outer interfaces of the
antibubble can take place faster. Such behavior would lead to nonlinearities
quickly, similar to how coatings on microbubbles have led to nonlinear
‘compression-only’ responses [45]. Another effect that likely plays a role is
resonance. Like microbubbles, antibubbles exhibit acoustic resonances [39],
and an antibubble driven near its acoustic resonance will oscillate more
strongly. In our microscopy experiments, the driving frequency of
$90.5\text{\,}\mathrm{kHz}$ corresponds to a resonant antibubble diameter on
the order of $100\text{\,}\mathrm{\SIUnitSymbolMicro m}$ [39]. The specific
resonant radius also depends on the core diameter, which can vary slightly
within a sample of antibubbles. However, this radius is clearly larger than
the antibubbles in our measurements, so one would expect that the larger
antibubbles would respond more strongly, and more nonlinearly, to the
ultrasound in our experiments, just as we observed.

Beyond the destructive processes of collapse or fragmentation, our
measurements also reveal a more gentle mechanism of release for the smaller
antibubbles. These bubbles do not significantly change shape but clearly empty
their core contents into the surrounding fluid, as indicated by the change in
transparency of the antibubble. The following dynamics are also characteristic
of microbubbles without a core, further supporting that they have released the
payload. The specific mechanism for gentle release remains unclear, but our
hypothesis is that small volumes of payload are ejected from the core by a
transient bridging of the inner and outer fluids, reminiscent of ‘kiss and
run’ exocytosis [46] in cellular systems. Such effects could be enhanced by
the fact that multiple droplets are encapsulated in the air bubble, so that
individual core droplets could be released without affecting others. Such
behavior could also play a role in the stepwise release observed in the
hydrophobic antibubbles, where some core contents are retained between pulses.
While our current measurement setup does not allow for high-speed fluorescence
measurements to confirm these behaviors, we speculate that the hydrophobic
shell stabilizes the structure in the presence of ultrasonic disruption, and
would quickly close any bridges between the inner and outer fluids, so that
only a small amount of the payload, or a single droplet from the multi-droplet
core, can be released at a given time. This mechanism, however, requires
further investigation.

##  3 Conclusion

We have demonstrated that antibubbles respond to low-intensity ultrasound by
releasing their payload in a controllable manner. By varying the antibubble
size and composition, the release pressure can be tuned within the 1-100
$\mathrm{kPa}$ range, and different release profiles can be achieved. A
nontrivial interplay of the antibubble composition and size leads to different
behaviors. For antibubbles with hydrophilic shells, the release pressure is
larger for smaller antibubbles, while the opposite is true for hydrophobic
antibubbles. Hydrophilic antibubbles displayed a characteristically sharp
pressure release threshold, and ejected the majority of the payload after a
single ultrasound pulse. By contrast, the smaller hydrophobic antibubbles
exhibited a unique stepwise release profile, making it possible to precisely
dose the payload delivery over time. A surprising result of our experiments is
that the antibubbles demonstrate robust low-intensity release behavior over a
wide range of frequencies ($<$$100\text{\,}\mathrm{kHz}$) to
($1\text{\,}\mathrm{MHz}$). Unlike conventional bubble-based release, which
requires intense, resonant excitation to rupture the bubble, the
responsiveness of antibubbles across a broad frequency range could be
advantageous in a wider range of applications. The low release threshold is
also specific to ultrasound, and the bubbles remain stable even in the
presence of mild mechanical agitation such as flows and stirring. While higher
background ultrasound intensities could spuriously trigger unspecific release,
targeted ultrasonic fields typically localize the ultrasonic pressure
sufficiently and therefore should not trigger release beyond the higher-
intensity focus. We have identified multiple distinct mechanisms that could
play a role in the release process under different conditions: nonlinear
oscillations, multi-bubble interactions, and transient bridging. Microscopy
data shows that these mechanisms can lead to a total release of the core
contents from hydrophilic antibubbles in some cases, while possibly leaving
partially-filled or empty microbubbles behind in others. It remains an open
question to identify if there are conditions that lead to total destruction of
an entire sample and complete gas dissolution. Further work is needed to
understand the specific role of each mechanism at different frequencies and
for specific antibubble compositions.

The ability to deliver payloads at low pressures, as well as the ability to
tune the delivery profile between single and stepwise release, make
antibubbles a unique carrier for payload delivery in diverse settings. Our
results lay the foundation for designing tailor-made formulations for specific
applications such as controlled drug delivery [47, 48], acoustic fabrication
[1], and healing materials [43]. More complex payloads including hydrophobic
drugs, enzymes and even simple organisms [49] can be encapsulated without
damage, supporting a rich set of delivery applications. In many of these cases
antibubble sizes similar to those studied in this work are directly relevant,
while others - primarily clinical - will require smaller antibubbles (e.g.
below $10\text{\,}\mathrm{\SIUnitSymbolMicro m}$). This motivates further
study of different shell compositions and investigations of different
encapsulation techniques to provide more control over sample sizes and
monodispersity. Further work in these directions could make it possible to
build a library of antibubbles whose response behavior can be selectively
addressed by pressure and frequency. Such a library would make it possible to
combine distinct antibubble formulations carrying different payloads, and
allow for specific triggering of single classes of antibubbles at a time,
opening the door to controlling complex biological or chemical processes using
antibubble carriers and low-intensity-ultrasound.

##  4 Experimental Section

###  4.1 Antibubble fabrication

The general fabrication of the different antibubbles was performed as
described elsewhere [33]. First, the inner aqueous solution was prepared by
dissolving in 3.44 g type I water (Millipore), 0.6 g NaOH 2 M, 0.3 g calcein
and 0.44 g maltodextrin. The final calcein concentration is around 119 mM.
Second, 0.8 g of fumed silica Aerosil R972 (Evonik) was dispersed in 16.4 g of
cyclooctane using an ultrasonic sonotrode (Hirscher, UP100) until no visible
fumed silica was observed, forming the intermediate oil phase. For the
external aqueous phase, 160 g of water, 20 g of maltodextrin, 20 g of mannitol
and 2 g of NaCl were combined. Then 100 g of this solution was used to
disperse either Aerosil R972 fumed silica alone or a mixture of Aerosil 200
with Aerosil R972 in a 2:1 mass ratio using rotor-stator homogenization
(Ultraturrax T18, IKA) and an ultrasonic sonotrode (Hirscher, UP100). The
final total fumed silica content of the outer phase is about 0.5 wt$\%$. Next,
4 g of the inner aqueous solution was dispersed in 17 g of the medium solution
using an ultrasonic sonotrode (Hielscher, UP100H, amplitude 100$\%$, cycle 1,
60 s) to form a W/O emulsion. Next, 5 g of the W/O emulsion was combined with
40 g of an external solution containing 0.5 wt$\%$ of fumed silica by rotor-
stator homogenization (Ultraturrax T18, IKA) to obtain W/O/W emulsions. The
mixture was first homogenized at a speed of 7000 rpm for 2 min and then 20 g
of sample was scooped out and transferred to a centrifuge tube. The remaining
sample was further homogenized at 14000 rpm and then transferred to another
centrifuge tube. Approximately 30 mL of the external solution without fumed
silica was added to each batch and then centrifuged at 500 RCF for 2 minutes.
The concentrate from the top was then transferred to a round bottom flask and
frozen at -60∘C for at least 8 h and then dried at -85∘C and 0.1 mBar for 24 h
(Martin Christ, Alpha 2-4 LSCplus). The resulting dry powder was stored and
used after reconstitution in aqueous NaCl 2$\%$.

Aerosil R972 and Aerosil 200 were generously provided by Evonik. Maltodextrin
2DE and 12DE were provided by Roquette. Sodium hydroxide, calcein,
cyclooctane, d-mannitol, and sodium chloride were purchased from Sigma-
Aldrich. All reagents were used without further purification.

###  4.2 Antibubble bursting

####  4.2.1 Fluorescence-pressure plots

For bulk fluorescence experiments, a lead zirconate titanate (PZT) disk
transducer with 25 mm diameter and 2 mm thickness (PI Ceramic) was glued to
the side wall of the cuvette using cyanoacrylate superglue (Loctite 401).
Electrical leads were soldered to opposite sides of the transducer and the
device was driven by a function generator (AFG2062, Tektronix) connected to a
power amplifier (ENI 2200L). A sample of antibubbles (10 mg powder
reconstituted in 4. mL of 3% NaCl solution) was filled into a 4 mL quartz
spectroscopy cuvette with an ultrasonic transducer bonded to one side. The
sample was illuminated using a fiber-coupled LED (Thorlabs M470F4), and
fluorescence was measured at 90 degrees using a fiber-coupled spectrometer
(Ocean Optics Maya 2000).

Acoustic pressures were measured using a needle hydrophone (Onda HNR-0500)
placed in the sound field. The voltage output from the hydrophone was
converted to a pressure value using calibration curves provided by the
manufacturer. Since standing waves are formed both in the cuvette and on the
glass plate, we scanned the pressure fields spatially to find and report the
peak pressures within an experiment.

The release thresholds are calculated for each shell composition based on at
least $n=3$ independent experiments where fluorescence was measured after
ultrasound pulses of increasing pressure. Between pulses, the sample was
gently shaken to redistribute the positively-buoyant antibubbles throughout
the cuvette. The release threshold is defined when the increase in normalized
fluorescence for a given pressure excitation is statistically significant
compared to the scatter in the low-pressure fluorescence data. We deemed an
increase significant when the fluorescence measurement was at least 2 standard
deviations above the average fluorescence for all lower pressures. The
threshold was then taken as the mean of the pressure where the fluorescence
increase was detected and the previous measurement pressure. The error bars in
the threshold are set by the spacing between pressure measurement points. As
shown in Fig. 2C and D, a response threshold could not always be determined.
In the case of Fig. 2C, a significant response was already observed at the
lowest excitation pressure tested. In the case of Fig. 2D, no response was
observed to the highest pressure measured.

####  4.2.2 Glass plate experiments

The glass plate excitation setup was designed to allow for ultrasound
excitation of the antibubbles while permitting transmission optical
measurements in a microscope. A PZT piezoelectric disk was glued to one face
of a $50\times 50$ mm glass slide and electrical leads were soldered to
opposing sides of the piezo. A sample chamber was created on the glass slide
using an adhesive Gene Frame (Thermo Fisher) with
$250\text{\,}\mathrm{\SIUnitSymbolMicro m}$ height. Antibubble samples were
suspended in a solution 3 wt.% NaCl and 0.3 wt% xantham gum and loaded into
the chamber. The chamber was sealed from above using a polyester transparency
film. The piezo was connected to a signal generator (Tektronix AFG1062) via a
53 dB power amplifier (ENI 2200L), and driven with a single,
$300\text{\,}\mathrm{\SIUnitSymbolMicro s}$ pulse of ultrasound at
$90.5\text{\,}\mathrm{kHz}$. At this frequency, the piezo excited plate waves
in the glass slide, which formed a standing wave pattern in the sample chamber
as shown in Figure S2. The peak pressure in the sample chamber was
$58\text{\,}\mathrm{kPa}$ ($MI=0.19$, peak intensity
$0.1\text{\,}\mathrm{W}\text{\,}{\mathrm{cm}}^{-2}$).

####  4.2.3 Microscopy characterization

Antibubble size distributions were measured using an inverted optical
microscope (Zeiss Observer D1) with a $5\times$ objective. Size distributions
are based on microscopy images, where the software ImageJ was used to measure
the diameter of antibubbles. The scale was determined using a calibration
slide.

Confocal microscopy was performed on a Zeiss LSM 900 (inverted) laser-scanning
confocal microscope. Brightfield and green fluorescence images of the entire
sample chamber were acquired as multiple tiles with a pixel size of
$12.5\text{\,}\mathrm{\SIUnitSymbolMicro m}$. The tiles were stitched together
in postprocessing after background correction. Images were acquired before and
after excitation with ultrasound. The excitation took place outside of the
confocal microscope and the sample was returned to the microscope for further
imaging.

High speed videos of antibubble responses to ultrasound were acquired at
120,000 frames per second using a Photron Nova S16 camera mounted onto an
Olympus BX61 upright microscope. Antibubble samples in the glass slide
excitation geometry were imaged using a $10\times$ objective, with a pixel
size of
$2\text{\,}\mathrm{\SIUnitSymbolMicro}\mathrm{m}\mathrm{/}\mathrm{p}\mathrm{x}$.
The high speed camera recording and ultrasound exposure were triggered
simultaneously via a TTL pulse. The camera recorded 50 frames before the
trigger and 49950 frames after, covering the entire pulse. The videos were
saved as an uncompressed .avi file and quantitative measurements were made
using the open source software FIJI [50].

###  4.3 Statistical Analysis

For the comparison of size distributions before and after ultrasound exposure
(Fig. S5) two samples of $n=300$ were compared using a Welch’s t-test, using
Origin software. Results were statistically different with a significance
level of $\alpha=0.05$ and p-value $P=2.539\times 10^{-5}$.

Supporting Information

Supporting Information is available from the Wiley Online Library or from the
author.

Acknowledgements

This work was supported by the European Research Council under the ERC
Advanced Grant Agreement HOLOMAN (No. 788296). The authors gratefully
acknowledge the data storage service SDS@hd supported by the Ministry of
Science, Research and the Arts Baden-Württemberg (MWK) and the German Research
Foundation (DFG) through grant INST 35/1314-1 FUGG and INST 35/1503-1 FUGG.

Supporting Information for: Antibubbles enable tunable payload release with
low-intensity ultrasound

Nicolas Moreno-Gomez* Athanasios G. Athanassiadis* Albert T. Poortinga Peer
Fischer

##  S1 Intensity and Mechanical Index Calculations

To facilitate more direct comparison with clinically-relevant ultrasound
metrics, here we report the mechanical indices and equivalent plane wave
intensities corresponding to the threshold pressure we identify in the main
text. The mechanical index is calculated as [5]

$MI=\frac{P}{\sqrt{f}},$

where $P$ is the peak negative pressure in $\mathrm{MPa}$ and $f$ is the
ultrasound frequency in $\mathrm{MHz}$.

While intensities are often used in a clinical context, the release phenomena
described here are driven by a pressure mechanism. The intensity in our
standing-wave-based excitation geometries is therefore zero. For more
meaningful comparison to clinical settings, we therefore calculate the
intensity of a plane wave with the same peak pressure as we applied in our
experiments. The intensity is therefore calculated as [5]

$I=\frac{P^{2}}{2\rho c},$

where $P$ is the peak pressure, $\rho$ is the fluid density, and $c$ is the
fluid sound speed. For the calculations, we assume the fluid medium is water
with a density of
$\rho=$1000\text{\,}\mathrm{k}\mathrm{g}\mathrm{/}\mathrm{m}^{3}$$ sound speed
of $c=$1500\text{\,}\mathrm{m}\text{\,}{\mathrm{s}}^{-1}$$.

Pressure | Frequency | Mechanical Index | Equivalent Plane Wave Intensity
---|---|---|---
($\mathrm{kPa}$) | ($\mathrm{kHz}$) | ($\mathrm{M}\mathrm{Pa}\mathrm{/}\sqrt{\mathrm{M}\mathrm{Hz}}$) | ($\mathrm{W}\text{\,}{\mathrm{cm}}^{-2}$)
7 | 1000 | 0.007 | 0.002
49 | 1000 | 0.049 | 0.080
88 | 1000 | 0.088 | 0.258
250 | 1000 | 0.250 | 2.083
58 | 90.5 | 0.19 | 0.11
Table S1: Mechanical Index and Equivalent Plane Wave Intensity corresponding
to the release thresholds and excitation pressures identified in the main
text.

##  S2 Supporting Figures

Figure S1: Calcein fluorescence in 0.3 M NaOH as function of Calcein
concentraion. Above the concentration of 3 mM the fluorescence intensity of
the calcein drops because of self-quenching of the dye. Figure S2: The
stability of rehydrated antibubbles was evaluated by monitoring their size
variation over a 20-hour time period. No noticeable alteration in diameter was
observed for any of the investigated formulations. A minimum of n = 190 data
points were assessed for each formulation.

(a) Aerosil 200 and Aerosil R972 - 7000 rpm

(b) Aerosil 200 and Aerosil R972 - 14000 rpm

(c) Aerosil R972 - 7000 rpm

(d) Aerosil R972 - 14000 rpm

Figure S3: Raw data for the bulk fluorescence measurements. Each panel shows
the recorded fluorescence intensity (arbitrary units) as a function of
pressure, reported as the peak hydrophone voltage (mVpp) measured in the
cuvette. Panel (a) and (b) correspond to the hydrophilic formulations, panel
(c) and (d) to the hydrophobic one (see main text). Peak-to-peak drive voltage
amplitudes were converted to peak acoustic pressures in the main text using a
calibration factor of $1.411\text{\,}\mathrm{kPa}\text{\,}{\mathrm{mV}}^{-1}$,
determined from hydrophone measurements. Different markers represent
independently measured samples. Figure S4: Size distribution for the
Hydrophilic sample homogenized at 7000 rpm. Figure S5: Measured pressure
amplitude at 90.5 kHz within the sample chamber on the glass slide.

##  S3 Supporting Videos

Video S1

Confocal Z-stack of a $\sim$$30\text{\,}\mathrm{\SIUnitSymbolMicro
m}$-diameter hydrophilic antibubble. Compared to Video S2, the encapsulated
calcein concentration was lower so that the core is visible. Background
fluorescence is not visible here because a lower exposure time and gain were
needed to observe the inner core.

Video S2

Confocal Z-stack of a $\sim$$30\text{\,}\mathrm{\SIUnitSymbolMicro
m}$-diameter hydrophobic antibubble. Because of the high concentration of
calcein loaded in the antibubble, the core is self-quenched and no
fluorescence is visible internally. Free calcein in the solution and bound to
free-floating silica is visible because of the long exposure and gain settings
on the microscope.

Video S3

Dynamics of 3 antibubbles during ultrasonic radiation at 90.5 kHz, recorded at
120000 fps.

Video S4

Dynamics of 2 antibubbles during ultrasonic radiation at 90.5 kHz, recorded at
120000 fps.

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