Fluorine in medical microbubbles – Methodologies implemented for engineering and investigating fluorocarbon-based microbubbles

Fluorine in medical microbubbles – Methodologies implemented for engineering and investigating fluorocarbon-based microbubbles

Accepted Manuscript Title: Fluorine in medical microbubbles–Methodologies implemented for engineering and investigating fluorocarbon-based microbubble...

948KB Sizes 0 Downloads 21 Views

Accepted Manuscript Title: Fluorine in medical microbubbles–Methodologies implemented for engineering and investigating fluorocarbon-based microbubbles Author: Marie Pierre Krafft PII: DOI: Reference:

S0022-1139(15)00058-5 http://dx.doi.org/doi:10.1016/j.jfluchem.2015.02.013 FLUOR 8519

To appear in:

FLUOR

Received date: Revised date: Accepted date:

25-11-2014 16-2-2015 22-2-2015

Please cite this article as: M.P. Krafft, Fluorine in medical microbubblesndashMethodologies implemented for engineering and investigating fluorocarbon-based microbubbles, Journal of Fluorine Chemistry (2015), http://dx.doi.org/10.1016/j.jfluchem.2015.02.013 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Ac

ce

pt

ed

M

an

us

cr

i

*Graphical Abstract - Pictogram

Page 1 of 31

*Graphical Abstract - Synopsis

Text for graphical abstract

Ac

ce pt

ed

M

an

us

cr

ip t

Fluorocarbon gases are a key element for the stabilisation of medical microbubbles currently used for ultrasound diagnosis, and with potential for molecular imaging and targeted drug and gene delivery. Fluorosurfactants that provide exceptional shell elasticity could uplift bubble technology. We review the principal laboratory microbubble preparation procedures and sizing methods, as well as techniques implemented for investigating Gibbs and Langmuir films, which are essential models for studying microbubble shell structure and properties.

Page 2 of 31

Highlights Fluorine in medical microbubbles – Methodologies implemented for engineering and investigating fluorocarbon-based microbubbles

cr

us

an

M



ed



ce pt



Microbubble contrast agents are being used in cardiovascular diagnostic procedures Microbubbles are also investigated for other diagnostic procedures, molecular imaging, targeted drug and gene delivery All commercial bubble contrast agents contain a fluorocarbon in their gas phase; fluorosurfactants could improve bubble technology The principal laboratory microbubble preparation procedures and sizing methods are reviewed and discussed Gibbs and Langmuir films are useful models for investigating microbubble shell properties

Ac

 

ip t

Marie Pierre Krafft

Page 3 of 31

1 MiniReview

Fluorine in medical microbubbles – Methodologies implemented for

cr

ip t

engineering and investigating fluorocarbon-based microbubbles

Marie Pierre Krafft

us

Institut Charles Sadron (CNRS UPR 22). University of Strasbourg, 23 rue du Loess, 67034

an

Strasbourg Cedex 2 (France).

*Corresponding author:

M

Dr. Marie Pierre KRAFFT Tel: (+33) 3 88 41 40 60 Fax: (+33) 3 88 40 41 99

Ac ce p

te

d

E-mail: [email protected]

Page 4 of 31

2 Abstract

Gaseous microbubbles are being used in cardiovascular diagnostic procedures. They have

ip t

further potential in diagnosis of tumours, vascular and blood flow abnormalities, as well as for targeted, ultrasound-triggered drug delivery and as intravascular mechanical devices. All

cr

the commercially available microbubble-based contrast agents comprise a fluorinated inner gas in their composition. Fluorinated components (fluorocarbon gases, self-assembling

us

perfluororoalkylated surfactants) can play a key role in the engineering, investigation and development of microbubbles. Fluorocarbon gases provide osmotic stabilization and a co-

an

surfactant contribution to interfacial tension reduction. Perfluoroalkylated surfactants selfassemble in aqueous media to form bubble shells with exceptional elasticity and resilience,

M

and, optionally, effective anchorage for surface functionalisation. This paper critically reviews the foremost methods used for preparing and monitoring microbubble suspensions.

d

In particular, it identifies some common pitfalls encountered in the determination of

te

microbubble sizes. The paper also reviews the methods used for collecting reliable data on the bubble morphology and shell structure that are required for understanding, controlling

Ac ce p

and improving their functional properties. Models for bubble shell studies include spontaneously adsorbed Gibbs films and spread Langmuir monolayers. Keywords: Fluorocarbon; Fluorinated surfactant; Microbubble, Self-assembled film; Ultrasound; Diagnosis, Targeted drug delivery.

Page 5 of 31

3

1. Introduction and scope Micrometer-size gas bubbles are being produced and investigated as agents for contrast ultrasound imaging, molecular imaging, targeted drug and gene delivery, and as

ip t

mechanical intravascular intervention devices [1-10]. When submitted to an ultrasound field, microbubbles alternately shrink and expand in response to the oscillations of the

cr

acoustic pressure caused by the incident sound wave. The volume expansion of the

us

microbubbles is maximal at a specific frequency referred to as the natural resonant frequency, which is inversely related to their size [11]. The microbubble resonance

an

produces a backscatter echo that allows their detection and localisation. The echogenicity (i.e. relative strength of the backscattered signal) is strongest near the microbubble’s resonance frequency. Microbubbles are being used primarily for the visualisation of the

M

cardiac chambers during echocardiography. Parenteral administration allows clear imaging of the vascular spaces, which “light up” when microbubbles are present. Microbubbles can

d

also help visualise liver lesions, vessel occlusions and abnormalities, tumours, and help

te

assess tissue perfusion. The concept of molecular imaging relies on the selective

Ac ce p

adherence of microbubbles bearing an appropriate ligand to a target receptor expressed by specific tissues. Several pathologies are being targeted, including angiogenesis, inflammation and atherosclerosis [1]. Further applications are being explored that include use as mechanical intravascular clot breaking devices [12] and as oxygen [13], gene and drug delivery systems [14, 15]. The latter application appears particularly versatile and promising: the appropriately modified microbubbles can be targeted to a specific receptor system and a drug cargo can then be released at the chosen location by triggering an ultrasound pulse of sufficiently high energy [15]. Drug or gene delivery can also be guided by external forces (e.g. a magnetic field by incorporating magnetic nanoparticles on the microbubbles [16]). Several commercial bubble products are currently available for diagnostic procedures, primarily to highlight the endocardial border. The products presently available or under

Page 6 of 31

4 development mainly comprise bubbles with polymeric shells (e.g. heat-denaturized albumin) and those self-assembled from molecular surfactants, mainly phospholipids.[15] All these bubbles include a fluorinated inner gas. FDA-licensed bubble products for

ip t

diagnosis of ventricular dysfunction involve SF6, C3F8, C4F10, C5F12, and C6F14. The properties of fluorocarbons that are used for this purpose include extremely low water

cr

solubility combined with high vapour pressure, extremely low surface tension, unexpected co-surfactant activity, and chemical and biological inertness [17-20]. physical

chemistry

of

self-assembled microbubbles

is

being

us

The

extensively

investigated, and in particular the properties of their shells that determine the microbubbles’

an

functional attributes such as size and stability control [8, 21-23], shell viscosity [24, 25], resonance frequency [26], capacity to generate sound harmonics [27-29], to enable gene

M

transfer [30, 31], and to undergo ultrasound-triggered disruption [32]. Progress over the existing products is desirable and includes increased stability, improved control of size and

d

size distribution, of response to ultrasound waves and other stimuli, ability to generate

te

ultrasound harmonics, and to undergo ultrasound-triggered disruption. Tuning the bubble’s shell properties for sonoporation, physiological barrier crossing, rupture for drug delivery,

Ac ce p

energy release for clot breaking are important biomedical goals. Rational microbubble engineering, control and in vivo monitoring, and assessment of

efficacy suppose improved understanding of the fundamentals that govern bubble stability and properties, and hence, the development of adequate practical preparation and investigation procedures. Introducing a fluorocarbon (FC) gas and/or a perfluoroalkyl chain (F-chain) in the structure of a shell-forming amphiphile, which enhances its propensity for self-assembly, strongly increases the stability of microbubbles [33]. The microbubble’s interfacial film (or shell) is essentially planar at the molecular level and alike to, and shares the structure and properties of monolayer surfactant films spread (Langmuir films) or spontaneously formed (Gibbs films) at the air/water interface. These films thus provide relevant models for investigating, understanding and controlling the properties of

Page 7 of 31

5 microbubbles. It has been shown that fluorinated components can strongly improve the stability of interfacial films and microbubbles and help control their properties [34, 35]. Recent studies of the monolayer behaviour of perfluoroalkylated (F-alkylated) amphiphiles

ip t

include those of highly fluorinated functionalized phospholipids [36], glycolipids [37], anchor-shaped bolaamphiphiles [38], and phosphates [25]. The case of semi-fluorinated

cr

alkanes (CnF2n+1CmH2m+1, FnHm diblocks), which are amphiphiles devoid of polar head is interesting. They form monolayers that are nanostructured and display regular arrays of

us

large surface micelles [19, 39]. The study of such patterned self-assembled systems might suggest new concepts in the design of surface-decorated microbubbles with tailored

an

properties.

It should be noted that there is a strong trend for most of the major industrial producers

M

of highly fluorinated compounds to replace long-F-chain (≥7 fluorinated carbons) compounds by shorter chain or more easily degradable chain (e.g. F-polyether chain)

d

compounds whenever possible, and this for environmental reasons [40-42]. Long-F-chain

te

compounds have indeed been found to disseminate, bioaccumulate and persist in the environment. Their production and use are now strictly regulated in the western countries.

Ac ce p

F-alkyl acids with F-chains having six or less fluorinated carbon atoms are not considered bioaccumulative. Numerous specific exemptions have however been granted for use of high performance long-F-chain compounds, especially for applications related to safety (e.g. aeronautics, metal plating procedures, therapeutics, medical devices) [42]. It should also be noted that the amount of fluorinated material required for a diagnostic test is minuscule. A typical effective dose for cardiovascular examination, although it involves ca. 108 bubbles, only involves micrograms of active injected material, meaning, for example, that one kg of F-butane allows performing on the order of one million diagnostic procedures. A blatant need has been recognized for reliable physicochemical data related to the understanding of F-chain-length dependence of colloidal and interfacial behaviour (e.g. self-

Page 8 of 31

6 association, surface film formation ability, viscoelasticity and other characteristics) relevant to emulsification, aerosol and bubble formation, adsorption, coating, and to effects on physiology, pharmacokinetics and environmental behaviour [43].

ip t

Where microbubbles are concerned, collection of valid physicochemical data requires first the production of well-defined, reproducible size-controlled microbubble test samples,

cr

and second, the implementation of appropriately selected physical methods, models and data treatments. It is the purpose of this paper to critically review the principal laboratory

us

methods used for preparing such samples and collecting such data. In particular, it aims at identifying common pitfalls, for example in the measurement of the microbubble sizes. The

an

paper is organised as follows: after some reminders on the requirements for intravascular administration (Section 2) and in vivo dynamics (Section 3) of microbubbles, we will present

M

the methods used in the laboratory for their preparation (Section 4). We will then describe the methods of investigation of microbubbles (Section 5), including for monitoring their

d

sizes and stability and determining the properties and structure of their surfactant shell.

te

2. Requirements for intravascular microbubble administration

Ac ce p

Most diagnostic ultrasound devices operate at frequencies in the 1-10 MHz range, which requires bubbles in the micrometer size range. Free passage through capillary beds supposes bubble sizes well below the size of the much more deformable red blood cells (∼7 µm). The target size is therefore in the 1 to 7 µm range. The lower size limit is still a matter of debate. The presence of large bubbles should be avoided even if the average diameter is adequate. The bubble size distribution should be narrow and devoid of a “tail“ of large bubbles. Improper physical parameters can result in untoward side effects [15]. Size and size distribution of particles are known to impact on biocompatibility and in vivo recognition, and hence, persistence in the blood circulation [44]. It is therefore essential that bubble sizes and size distributions be properly controlled. The bubble shell should not impede the bubble’s ability to resonate when exposed to ultrasound waves. Therefore, the bubbles

Page 9 of 31

7 must have a highly flexible, elastic soft shell. They must be stable enough to resist repeated cycling through the heart and lungs. They must be provided in a form that allows transportation and storage for practical use in hospitals and in the radiologist’s office. Their

product must allow consistent dosing and should be user-friendly.

ip t

manufacture needs to be highly reproducible and cost effective. Finally, the injectable

cr

Of course, the product’s toxicity and side effects must be minimal. The amount of active components involved in a diagnostic dose is generally orders of magnitude below their

us

lethal dose. On the other hand, improper characteristics (e.g. bubble sizes, excessive polydispersity) can elicit side-effects. The incidence of side effects of microbubble contrast

an

agents, including hypersensitivity, allergic reactions, myocardial side effects, headaches, abdominal pain and temporary embolisms, appears to be much lower than that observed

M

for computed tomography (CT) and magnetic resonance imaging (MRI) contrast agents [45]. However, damage to blood vessels at the microvascular level is possible following

te

times.

d

microbubble destruction when using high microbubble concentrations and long insonation

Ac ce p

3. In vivo bubble dynamics

Once injected in the blood circulation, microbubbles are submitted to several forces that

concur to foster their dissolution in the surrounding fluid (Fig. 1). All bubbles are submitted to Laplace pressure, PL = 2γ/r, where γ is the interfacial tension on the bubble’s surface and r the bubble’s radius. As bubbles shrink, the Laplace pressure increases, which accelerates gas dissolution and further bubble shrinkage. Arterial blood pressure exercises another effective strain on bubbles and amounts to roughly 100 torr. Exposure to ultrasound energy submits the bubble to mechanical stress. All these forces combine to drive rapid diffusion of the standard gases (air, nitrogen) in water and consequent deflation/dissolution of an air bubble in the surrounding aqueous medium. Metabolism of the oxygen present in a circulating air bubble also contributes to bubble shrinking. When a FC gas is present it is

Page 10 of 31

8 progressively excreted by expiration when traversing the lung [44], thus reducing its stabilization effect. In vitro microbubble deflation was shown to involve expulsion of fragments of phospholipid bilayer through complex collapse and shedding mechanisms

an

us

cr

ip t

[46].

M

Figure 1. Representation of the forces that are exerted on a microbubble in the blood circulation during ultrasound examination and determine their dynamics, and of the forces that can be implemented to counteract the former.

d

In order to achieve sufficient microbubble stability and intravascular persistence for

te

effective practical use, it is necessary to counteract these phenomena. Effective

Ac ce p

stabilization can be gained by reducing inner gas solubility, reducing interfacial tension, optimizing bubble shell resistance to bubble size reduction, and hence, control its viscoelastic properties.

4. Laboratory microbubble preparation procedures The standard procedures available for producing microbubbles in the laboratory include

sonication and fractionation, electrohydrodynamic atomisation (EHDA) and microfluidic techniques. 4.1. Sonication or mechanical agitation followed by fractionation Reliable physicochemical studies necessitate the availability of well-defined, narrowly dispersed reproducible populations of microbubbles of predetermined sizes. Sonication and mechanical agitation allow the rapid production of milliliter- to liter-size samples containing

Page 11 of 31

9 large numbers of microbubbles. These samples suffer, however from high polydispersity (polydispersity index, defined as the ratio between the standard deviation and mean diameter in percentage, PDI > 30%) and are therefore not optimal for medical applications.

ip t

Much less polydisperse populations of bubbles can be obtained from these preparations by subsequent fractionation using flotation or centrifugation. A good correlation was found

cr

between measured bubble size values and those predicted by a dynamic model developed for the size fractionation by flotation of microbubbles having an albumin shell [47]. A fraction

us

of submicrometric microbubbles (∼0.45 µm), with a shell made of Span and Tween surfactants, has been separated by centrifugation or gravity from a larger mean-sized

an

sonicated preparation) [48].

We have shown that a polydisperse preparation of dimyristoylphosphatidylcholine

M

(DMPC)-coated microbubbles stabilized by F-hexane and produced by sonication could be fractionated simply under the action of gravity [21]. Narrowly dispersed populations having

d

mean radii of 1.6 and 5.4 µm have thus been isolated (Fig. 2). Centrifugal fractionation of

te

microbubbles coated with distearoylphosphatidylcholine (DSPC) and stabilized by F-butane

Ac ce p

has also proved successful [49].

Figure 2. Preparation of microbubble samples with controlled size and size distribution. Sonication of an aqueous dispersion of the surfactant is achieved under an atmosphere of air saturated with Fhexane. After dilution and washing of the bubbles, fractionation is achieved by flotation or centrifugation. In the former case, sampling at different precise levels in a graduated tube provides populations of bubbles with predetermined sizes [21, 22].

Page 12 of 31

10 Reproducibility of sonicated preparations requires considerable attention, including in the slightest details: size and shape of the vessel; liquid level; overlying atmosphere (usually FC-saturated nitrogen); sonication time, frequency and energy; probe size and tip

ip t

shape; position of sonicating probe with respect to the surface of the liquid, temperature, etc. The subsequent washing, dilution, flotation or centrifugation operations also require

cr

precise timing, temperature control and experimental know-how. Use of flotation for separation of differently sized bubble preparations involves sampling at different levels in a

us

given graduated tube and readily provides populations of bubbles with predetermined size. 4.2. Coaxial electrohydrodynamic atomization and microfluidics

an

Significant progress in terms of reducing polydispersity has been achieved using coaxial electrohydrodynamic atomization (CEDHA) and microfluidic techniques [50-52]. Bubbles

M

between 2 and 8 µm in diameter with very low polydispersity (PDI < 2%) have been obtained by CEDHA [53]. The number of microbubbles formed is smaller than that obtained

d

using sonication, but the production is continuous. Microfluidic technology offers a high

te

level of control over size distribution. Monodisperse microbubbles down to 2 µm have been obtained with a flow-focusing microfluidic technology [54]. These techniques are generally

Ac ce p

slow and limited to small samples. The production of such small bubbles with microfluidics requires very fine channels and high gas flow rates, which augments polydispersity [51, 52]. Another drawback appears when preparing microbubbles smaller than 10 µm: the tendency for microchannels to clog.

Commercial microbubbles are usually rather polydisperse (some microbubbles are as

large as 32 µm) [55] and they are not distributed in a ready-for-use form. They need generally to be reconstituted in the radiologist’s suite, for example by mixing a spray dried or lyophilized powder with an aqueous solution.

5. Microbubble investigation methods Monitoring

and

controlling

microbubble

production,

matching

specifications,

characterizing products and gaining information on the structure and physical properties of

Page 13 of 31

11 bubble shells, including via model interfacial films, require implementation of a range of appropriately chosen experimental methods. When fluorinated surfactants are used, investigation of the interfacial films is often facilitated by the huge difference in electron

contrast in electron microscopy and scattering profiles.

19

ip t

density of fluorinated versus hydrogenated sub-layers. This translates into enhanced F provides an additional sensitive

cr

probe for NMR studies. 5.1. Sizing and stability monitoring

us

The methods most commonly used for determination of size distributions in microbubble suspensions are the electrical zone sensing (EZS), the single-particle optical

an

sensing (SPOS), the static (also known as laser diffraction) and dynamic (also called photon correlation spectroscopy) light scattering methods. More recently, a multifrequency

M

acoustical attenuation method (MFA) was developed specifically for investigation and monitoring of microbubbles. The first two methods are based on devices that count the

d

bubbles by analysing the signals generated by individual particles, whereas the third and

te

fourth methods use diffractometers that analyze the light scattered by a collection of bubbles. The acoustical method also addresses collections of bubbles. The measurements

Ac ce p

performed with counters require accumulation of data collected from a large number of individual bubbles in order to provide statistically sound size distributions. Precision and reliability depend on sampling issues, dilutions, stirring, and involve calculation approximations (usually embedded in the data treatment software). The most important complications encountered in accurate bubble sizing arise from the fact that the bubbles tend to float due to their buoyancy, and that the test sample is modified during the measuring process, which thus prevents monitoring of the stability of a given bubble sample over time. Other limitations of these methods and of the corresponding commercial instruments are that, with the exception of the MFA method, they have been devised for other purposes, meaning that use for bubble sizing requires caution.

Page 14 of 31

12 5.1.1. Electrical zone sensing method Electrical zone sensing (EZS) was initially developed for counting blood cells and is the most frequently used method for microbubble sizing [56]. Microbubbles suspended in

ip t

an electrolyte solution are channelled through a small aperture positioned between two electrodes. The voltage applied across the aperture creates a "sensing zone". As bubbles

cr

pass through this sensing zone, they displace their own volume of electrolyte, momentarily increasing the impedance of the aperture. The change in impedance produces a pulse that

us

is digitally processed in real time. The height of the pulse is proportional to the volume of displaced electrolyte solution. The volume of the bubble is then approximated from the

an

volume of displaced solution using a model for conversion into a spherical object. Analysis of numerous pulses enables a size distribution to be acquired and displayed in bubble

M

volume and diameter. Several thousand bubbles per second are individually counted and sized. The size range accessible to this method spans from ~0.4 to ~1200 µm, depending

d

on the diameter of the aperture (from 15 to 2000 µm). This method is independent of

te

microbubble refractive index and density. In addition, a metering device is used that draws a known volume of bubble suspension through the aperture; a count of the number of

Ac ce p

pulses yields the concentration of bubbles in the sample. However, a first drawback of this method is related to the data treatment model, which is valid only when the relative sizes of the bubbles and aperture are correctly dimensioned. It is usually accepted that the bubble should be smaller than the aperture by 60% in order to avoid significant edge effects. On the lower size side, the bubble needs to be larger than the aperture by 10% for the model to be valid. Approximations are needed for particles outside this range. Also, the Coulter Counter has been developed for red blood cells, which have a tendency to sediment. Therefore, the geometry of the measuring cell and stirring device were designed to redisperse dense red blood cells. When microbubbles are measured, this cell design strongly favours buoyancy, which is detrimental to accuracy. Finally, it is not possible to monitor the stability of microbubbles over time using EZS, as the sample is altered by the method.

Page 15 of 31

13 5.1.2. Optical methods The single-particle optical sensing (SPOS) method, also called optical particle counting (OPC), has originally been developed to analyze contamination of fluids by

ip t

particles. The suspension of microbubbles is drawn through a small "photozone", which is a narrow, uniformly lit slab-like region [57]. The bubble suspension must be sufficiently dilute

cr

so that the bubbles pass one by one through the illuminated region. The passage of a bubble through the sensing zone causes light diffraction, with a magnitude that depends on

us

bubble size. Detection can be achieved by light scattering or obscuration. One serious limitation of this method is due to the low concentration of bubbles that is required. This

an

makes sampling delicate as it can eliminate the larger microbubbles, which are subject to rapid buoyancy. In the case of light obscuration detection, a further limitation is that the

M

model used for data treatment is only valid if the bubble is larger than the wavelength of the light by a factor of at least five in order to avoid diffraction (in which case it is necessary to

d

calculate the form factor). This strongly limits the use of this technique for gas bubble

te

analysis. It can be improved by using multi-angle light scattering detection, provided the bubble is larger than half the wavelength of the light, but this detection mode is seldom

Ac ce p

used. As for EZS, bubble buoyancy limits the use of SPOS, and likewise, it is not possible to analyse the stability of a given bubble sample over time. Optical microscopy can also be used to determine bubble size distributions. Sampling

issues and interferences of microbubbles with the plate wells often compromise the accuracy and reliability of the measurements. One further serious weakness of the method is that microbubbles smaller than ∼1 µm are not detectable. 5.1.3. Light scattering Static light scattering (SLS) is used to extract bubble size distributions from a light scattering pattern based on the theories of Mie and Fraunhaufer. This method is rapid and allows for larger bubble concentrations than counters (EZS or SPOS) [56]. As noted above, the range of sizes covered by SLS is well adapted to medical microbubbles. The lower limit

Page 16 of 31

14 is about 0.5 µm. However, the standard commercial measuring cell (which is high and narrow) and agitation device have usually been designed to re-disperse particles that tend to sediment, not particles that tend to float. Monitoring bubble stability over time is therefore

ip t

difficult. Dynamic light scattering (DLS) or photon correlation spectroscopy (PCS) is based on

cr

the time-dependent fluctuations of the intensity of the scattered light provoked by the Brownian motion of suspended particles. It addresses particle sizes ranging from a few

us

nanometres to about 100 nm, that is, well below the size of the bubbles investigated for biomedical uses. Furthermore, bubble buoyancy significantly affects Brownian motion,

an

while stirring is, in this case, prohibited. DLS actually measures correlation curves and fits them to an arbitrary chosen model, which allows estimation of the diffusion coefficient of the

M

particles, from which the hydrodynamic volume is calculated using the Stoke-Eisenstein equation. This method is therefore not appropriate for measuring distributions of micron-

d

sized bubbles [56].

te

5.1.4. Acoustic attenuation measurements The multi-frequency acoustical (MFA) method is based on the measurement of the

Ac ce p

attenuation coefficient of an ultrasound wave that propagates through a dispersion of bubbles. Its principle is schematically depicted on Figure 3 [21]. The microbubble dispersion is injected in a glass cell filled with an aqueous solution through which an ultrasound wave propagates. The decay of the intensity of this wave after having traversed the test sample is monitored at multiple frequencies. The sound attenuation coefficient α is maximal when the bubble’s resonance frequency is equal to the ultrasound excitation frequency. Since the bubble resonance frequency f is proportional to the bubble’s radius r, we can extract the mean size and size distribution of the microbubbles from the decay curves [21, 22, 58]. The method is particularly well adapted to monitoring soft-shell microbubble size and size distribution as a function of time, determining bubble stability and

Page 17 of 31

15 half-lives, providing information on deflation mechanism, assessing the effects of internal

an

us

cr

ip t

gases (e.g. a FC).

Figure 3. Microbubble sizing using an ultrasound attenuation method. a) the measuring cell in which

M

the bubble aqueous dispersion is insonated; b) acoustic attenuation coefficient versus ultrasound frequency; c) some of the equations used in the data treatment.

d

Major assets include the use of low-power ultrasound waves, thus minimizing the impact

te

of the sound wave on bubble characteristics, and a very precise measurement of sound absorption coefficients (Fig. 3). The MFA method presents definite advantages over the

Ac ce p

above methods. First, the measurement is independent of bubble movement and rapid (less than 0.5 s). The sample can therefore be stirred in order to effectively avoid bubble floating and aggregation. Second, the measurement is performed on the whole population. Third, the cubic geometry of the cell and its large volume (~140 mL) reduce the effect of the walls and facilitate homogenous agitation. Fourth, data treatment involves a minimal amount of approximations and modelling. Fifth, the measurement does not alter the sample, allowing for monitoring of bubble size, and, hence, stability upon time. Six, the device is fully automated, with the possibility of collecting many hundreds of data points over long time periods (~e.g. 8 h). Other acoustical cells [59, 60], including cells with only one transducer that plays both the roles of the emitter and of the receiver, as is the case for medical echographs [61-63], have been used for measuring the attenuation coefficient of microbubbles. These methods use three or four broadband emitting transducers to cover

Page 18 of 31

16 the appropriate frequency range (2 – 30 MHz). In our case, seven narrowband signals with different frequencies covering the whole frequency spectrum are being emitted by one single transducer. The main advantage of processing numerous narrowband signals, rather

ip t

than a smaller number of broadband signals is a much higher precision of the Fourier transform of the signal, owing to a better signal/noise ratio in the emitting range. The

cr

precision of the attenuation coefficient is, therefore, significantly improved. The advantage of using one emitting transducer and one receiving transducer instead of one that plays

us

both roles is that the signal has to cross only once the zone of stirring that inevitably creates perturbations.

an

The MFA method was validated repeatedly by concurrent results with optical microscopy and static light scattering, three complementary methods that we routinely used for

Ac ce p

te

d

M

determining microbubble size and size distributions (Fig. 4) [21].

Figure 4. Comparison of bubble size and size distribution measurements in a dispersion of Fhexane-stabilized

microbubbles

of

dimyristoylphosphatidylcholine

using

three

independent

microbubble sizing methods: optical microscopy (bars histograms), static light scattering (grey lines), and multi-frequency acoustical attenuation (black circles) [21]. The two populations of microbubbles (mean radius of ~1.5 µm (a) and ~5.0 µm (b)) were fractioned by flotation.

5.2. Gibbs and Langmuir films: privileged models for studying bubble shell properties Two-dimensional monomolecular Gibbs and Langmuir films made from bubble components provide valuable complementary models for the investigation of bubble shells in the presence or absence of fluorinated components, as the curvature of the shell is essentially zero at the molecular scale. Gibbs films form spontaneously when surfactants

Page 19 of 31

17 present in a solution adsorb at interfaces. They can be conveniently investigated by bubble profile analysis tensiometry (Fig. 5). The shape of a gas bubble immersed in a liquid is determined by interfacial tension and buoyancy. For bubble profile analysis, a microlitre-

ip t

size rising bubble is formed at the tip of a steel or Teflon capillary in a surfactant solution or dispersion. An effective instrument (Tracker®) is commercialised by Teclis (Longessaigne,

cr

France). This type of tensiometry is ideal for monitoring the time-dependence of interfacial

M

an

us

tensions and the surfactant’s adsorption kinetics at an interface [64].

Figure 5. Dynamic bubble profile analysis tensiometry performed on a millimetric bubble allows

d

determination of the rate of adsorption of a surfactant as a Gibbs film and the interfacial tension

te

reduction caused by bubble components. The bubble can be static or submitted to controlled surface variations by applying sinusoidal oscillations of various frequencies.

Ac ce p

The bubble can also be submitted to periodical surface variations through sinusoidal oscillations, a procedure that was found to substantially accelerate the adsorption process and allow investigation of surfactant systems that are too slow to reach equilibrium in standard conditions [65]. Viscoelastic

properties

of

thin

surfactant films

at

interfaces

are

preeminent

characteristics that often determine practicability of self-assembled colloids (emulsions, microbubbles, foams) for a wide range of applications [66]. They are particularly important for determining bubble resilience under ultra-sound and other strains. Dilational rheology allows measurement of the interfacial tension response to dilational/compression stresses (sinusoidal oscillations) applied to a surfactant film adsorbed from a solution to the surface of an oscillating air bubble. It has recently been used to study soluble surfactants [67].

Page 20 of 31

18 Dilational rheology enables determination of kinetic processes and thermodynamic parameters, and is complementary to shear rheology; both types of deformation modes exist in most practical situations. The dilational viscoelastic modulus, E = ∆γ/∆A, is defined

ip t

as the ratio of the variation of the interfacial tension ∆γ to the relative variation of the bubble’s surface area ∆A. E is a complex quantity that depends both on subtle changes in

cr

the adsorbed layer’s structure and on the frequency of the oscillatory perturbation.

us

Langmuir film investigation is achieved by depositing a surfactant solution, usually in a volatile organic solvent, on the surface of water enclosed in a Teflon trough and limited by

te

d

M

an

mobile Teflon barriers (Fig. 6).

Ac ce p

Figure 6. Langmuir monolayer. a) An organic solution of a water-insoluble surfactant is spread as a monolayer at the surface of water in a trough fitted with barriers that allow compression of the film while surface pressure is monitored. b) The surface pressure versus molecular area isotherm reveals the successive phase transitions undergone by the monolayer, until it collapses. c) Domains of liquid-condensed phase coexisting with a continuous liquid-expanded phase, as visualized by fluorescence microscopy.

Moving the barriers while measuring the pressure (as with a Wilhelmy plate) allows compression (or expansion) of the film on a given sub-phase and recording of surface pressure versus molecular area isotherms. It allows determination of collapse pressure, which reflects film stability. Langmuir film compression is particularly effective for assessing phase structure and phase transitions of the monolayer. The phases encountered as pressure increases are categorised as gaseous (G), liquid expanded (LE), liquid condensed (LC), and solid (S). Examples of isotherms are shown in Figure 7 [68].

Page 21 of 31

cr

ip t

19

us

Figure 7. Compression isotherms of Langmuir monolayers of a) [C6F13(CH2)2]2P(O)N(CH2CH2)2O; b) [C8F17(CH2)2]2P(O)N(CH2CH2)2O and c) [C9F19CH2]2P(O)N(CH2CH2)2O surfactants, illustrating markedly different phase behaviours. The monolayer of a) is in a liquid expanded state throughout

an

compression, whilst that of c) is in a liquid condensed state. The monolayer of b) exhibits a transition from a liquid expanded to a liquid condensed phase [68].

M

The Langmuir trough can be protected against evaporation, allows control of overlying atmosphere, and can be fitted with diverse observation instruments, such as for

d

fluorescence and Brewster microscopies, grazing incidence X-ray diffraction (GIXD),

te

grazing incidence small-angle X-ray diffraction (GISAXS) or X-ray reflectivity. The Langmuir-Blodgett technique allows lifting of monolayers from a water surface at any given

Ac ce p

surface pressure, and their subsequent deposition on solid substrates for further investigation (Fig. 8), for example by atomic force microscopy (AFM).

Figure 8. a) Langmuir-Blodgett transfer of a monolayer from the surface of water onto a solid substrate (silicon wafer, mica, etc.) allowing further investigation, for example by atomic force microscopy b).

Page 22 of 31

20 5.3. Bubble dissolution and shell morphology – Optical methods It is important to determine the fate of a microbubble suddenly suspended in a liquid medium containing different gases, a situation encountered during the injection of a

ip t

microbubble ultrasound contrast agent into the bloodstream. An experimental system has been developed which isolates the microbubbles in a permeable hollow fiber submerged in

cr

a perfusion chamber that allows rapid exchange of the external aqueous medium [69]. Results obtained on SF6 microbubbles coated with phospholipids showed that the growth

us

regime was less pronounced than when sodium dodecyl sulfate was used as the shell component, and three dissolution regimes were identified, including rapid dissolution back

an

to the original diameter, steady dissolution with a nearly constant wall velocity, and stabilization near ∼10 µm diameter.

M

The mechanical properties of the microbubbles’ shell play an important role in the stability of microbubbles. The observation of their shell morphology can provide valuable

d

information on these properties. In practice, monitoring the deflation of small, fast moving 1

te

– 10 µm bubbles in an aqueous dispersion using optical microscopy is difficult. Therefore experimental setups have been designed for studying larger bubbles. A simple method has

Ac ce p

recently been devised for visualizing the surfactant shell of bubbles of ~20-100 µm during growth or deflation under the effect of temperature [25]. It consists in generating the bubbles directly on the observation stage of the microscope by controlled heating of the surfactant solution. Conversely, their deflation can be monitored during cooling. Fluorescence microscopy (FM) was used to monitor the influence of FC gases on the

LE/LC transition of DPPC in Langmuir monolayers (Fig. 9) [20, 70]. The fluorescent dye used dissolves preferentially in the disordered LE regions of the monolayer, which therefore appear bright while the LC regions remain dark. The method allows distinction among phases, delineation of organized domains, determination of phase transitions, monitoring of transformation, etc. FM was also carried out directly on large bubbles (hundreds of µm) to

Page 23 of 31

21 visualise the LE/LC phase transitions [71, 72], and allowed demonstration of the effect of

cr

ip t

structure on permeation of a monolayer by gases.

us

Figure 9. Fluorescence microscopy showing the effect of F-octyl bromide gas on the occurrence of the liquid condensed/liquid expanded phase transition in a Langmuir monolayer of DPPC.

an

FM also demonstrated the effect of a partially fluorinated alcohol on the organisation of a DPPC monolayer. It showed that the F-alcohol acts as a lineactant at the periphery of the

M

condensed phospholipid domains and modifies their shape and organisation [73]. These results provide valuable information for the design of new lung surfactant substitutes.

d

Brewster angle microscopy (BAM) is based on the fact that no light is reflected when a

te

light beam linearly polarized parallel to the plane of incidence hits the air/water interface at an angle of 53.15° (an angle that is determined by the relative refractive indices of air and

Ac ce p

water, Fig. 10).

Figure 10. Brewster angle microscopy of a Langmuir monolayer. a) No light reflection occurs on pure water at the Brewster angle; b) light is reflected when an interfacial surfactant film is present. c) Brewster angle micrograph of a film of C10F21(CH2)OP(O)(OH).1Na spread on water, showing a pattern of micron-size self-assembled hexagon-shaped domains [74].

Page 24 of 31

22 The conditions for extinction are no longer satisfied when a monolayer is present at the interface. The monolayer appears brighter than water, which remains dark. The intensity of the reflected light is proportional to the film’s thickness, which can be

ip t

quantitatively determined in some cases. Figure 10 provides an example of BAM micrograph showing microdomains formed within a Langmuir film of the sodium salt of an

cr

F-alkyl phosphate surfactant [74]. 5.4. Bubble shell structure – Scattering methods

us

The structure and properties of molecular layers formed at the air/water interface are known to depend on a delicate balance between the polar head(s) and hydrophobic

an

chain(s) of the amphiphiles that make them up. The main driving forces for monolayer formation are the repulsion between heads, which allows the spreading of the amphiphiles,

M

and the hydrophobicity of the chains, which prevents molecules from dissolving in the aqueous phase [75]. Compression isotherms, combined with grazing incidence X-ray

d

diffraction (GIXD), allow determination of the influence of the different segments of the

te

amphiphiles on the organization of the thin films [76-78]. GIXD can assess the interactions between the surfactant molecules and their degree of ordering, and determine lattice

Ac ce p

parameters for crystalline surfactant films. This can be achieved either directly on water or after transfer on a solid substrate. GISAXS, that is, GIXD conducted at smaller angles, can assess the presence of domains (tens of nanometres in size) that are regularly organized on water. X-ray reflectivity measurements allow determination of electron density across a thin film on the water surface or after deposition on a solid substrate. A fluorinated sublayer is readily differentiated from a hydrogenated sub-layer and their thickness can be measured, as well as the orientation of fluorocarbon and hydrocarbon chains relative to the film’s surface (Fig. 11) [79].

Page 25 of 31

cr

ip t

23

us

Figure 11. X-ray reflectivity measurements on a monomolecular film of C8F17C16H33 (F8H16) diblock transferred on a silicon wafer. The heights of the C8F17 and C16H33 sub-layers were determined to be 1.0 nm and 1.9 nm, respectively (the fully extended length of the diblock is 3.3 nm). The fluorinated

6. Conclusions and perspectives

an

and hydrogenated blocs are pointing toward air and silicon wafer, respectively [79].

M

Careful implementation of the above outlined preparation and investigation methods have improved our control and understanding of microbubble formation, stability and

d

dissolution, structure and properties, and have allowed precise assessment and control of

te

practical issues related to their use in medicine. The Reader may find examples of advances gained through use of these methods in our

Ac ce p

Laboratory concerning the contributions of perfluoroalkylated components (fluorocarbon gases and fluorosurfactants) towards microbubble engineering and function, including understanding and implementation of FC gas-driven osmotic bubble stabilization [35]; recognition that the actual stabilization effect is orders of magnitude larger than predicted [35]; demonstration of a significant co-surfactant effect of a bubble’s internal FC gas with phospholipids, resulting in a lowering of the interfacial tension [80]; production of small bubbles that, contrary to common credence are more stable than larger ones of same composition [21]; evidence that the inner FC gas can increase the collapse pressure of a phospholipid monolayer and enhance its compressibility and fluidity [35]; establish that a FC gas can drastically modify, and actually reverse the outcome of the competitive adsorption of a phospholipid (DPPC) versus a protein (bovine serum albumin) at the

Page 26 of 31

24 air/water interface [81]; establish that some fluorosurfactants can provide “super” elastic bubble shells [25]; demonstrate pH control of microbubble formation, stability and properties using perfluoroalkyl phosphate surfactants [25]; monitoring and ascertainment of

ip t

microbubble surface decoration and functionalization, as by grafting superparamagnetic iron nanoparticles on their surface [25, 82].

cr

The knowledge gained through studies of fluorocarbon effects on microbubbles and related interfacial surfactant films is essential for microbubble and interface science and

us

technology understanding and engineering. This knowledge is also relevant to the design and control of other self-assembled colloidal systems, including vesicles [83], emulsions

an

[84] and foams [85]. It is also germane to other situations in which fluorocarbons or fluorocarbon chains come in contact with biological membranes, as in potential treatment of

M

lung surfactant deficiencies and respiratory conditions [20, 70, 86], preservation of organs and cells [87, 88], and control of cell aggregation [87].

d

Acknowledgements

te

The author thanks the European Commission (FP7 2007-2013; grant n° NMP3-SL-2008214032), the French National Research Agency (ANR, grant n° 2010-BLAN-0816-01), and

Ac ce p

the GIS Fluor for financial support. Teclis (Longessaigne, France) is acknowledged for technical help.

7. References

[1] G.M. Lanza, S.A. Wickline, Progr. Cardiovasc. Dis., 44 (2001) 13-31. [2] E.S. Schutt, D.H. Klein, R.M. Mattrey, J.G. Riess, Angew. Chem. Int. Ed., 42 (2003) 3218-3235.

[3] J.G. Riess, Curr. Opin. Colloid Interf. Sci., 8 (2003) 259-266. [4] J.R. Lindner, Nature Rev. Drug Disc., 3 (2004) 527-532. [5] E.C. Unger, T. Porter, W. Culp, R. Labell, T. Matsunaga, R. Zutshi, Adv. Drug Deliv. Rev., 56 (2004) 1291-1314. [6] A.L. Klibanov, Invest. Radiol., 41 (2005) 354-362. [7] S. Hernot, A.L. Klibanov, Adv. Drug Deliv. Rev., 60 (2008) 1153-1166. [8] S. Sirsi, M. Borden, Bubble Sci. Eng. Technol., 1 (2009) 3-17. [9] K.W. Ferrara, M.A. Borden, H. Zhang, Acc. Chem. Res., 42 (2009) 881-892.

Page 27 of 31

25 [10] M. Aryal, C.D. Arvanitis, P.M. Alexander, N. McDannold, Adv. Drug Delivery Rev., 72 (2014) 94-109. [11] H. Medwin, Ultrasonics, 15 (1977) 7-13. [12] A. Maxwell, C. Cain, A. Duryea, L. Yuan, H. Gurm, Z. Xu, Ultrasound Med. Biol., 35

ip t

(2009) 1982-1994. [13] J.J. Kwan, M. Kaya, M.A. Borden, P.A. Dayton, Theranostics, 2(2012) 1175-1184. [14] J.J. Rychak, A.L. Klibanov, Adv. Drug Delivery Rev., 72 (2014) 82-93.

cr

[15] E. Unger, T. Porter, J. Lindner, P. Grayburn, Adv. Drug Delivery Rev., 72 (2014) 110126.

us

[16] D. Vlaskou, P. Pradhan, C. Bergemann, A.L. Klibanov, K. Hensel, G. Schmitz, C. Plank, O. Mykhaylyk, AIP Conf. Proc., 1311 (2010) 485-494.

[17] J.G. Riess, M.P. Krafft, Mater. Res. Soc. Bull., 24 (1999) 42-48.

an

[18] M.P. Krafft, J.G. Riess, Perfluorochemical-based oxygen therapeutics, contrast agents, and beyond, in: A. Tressaud, G. Haufe (Eds.) Advances in Fluorine Science - Fluorine and Health. Molecular Imaging, Biomedical Materials and Pharmaceuticals, Elsevier,

M

Amsterdam, 2008, pp. Chapter 11, 447-486.

[19] M.P. Krafft, J.G. Riess, Chem. Rev., 109 (2009) 1714-1792.

(2006) 3184-3192.

d

[20] F. Gerber, M.P. Krafft, T.F. Vandamme, M. Goldmann, P. Fontaine, Biophys. J., 90

te

[21] S. Rossi, G. Waton, M.P. Krafft, ChemPhysChem, 9 (2008) 1982-1985. [22] S. Rossi, G. Waton, M.P. Krafft, Langmuir, 26 (2010) 1649-1655.

Ac ce p

[23] S. Dicker, M. Mleczko, G. Schmitz, S.P. Wrenn, Ultrasonics, 53 (2013) 1363-1367. [24] N.A. Hosny, G. Mohamedi, P. Rademeyer, J. Owen, Y. Wu, M.-X. Tang, R.J. Eckersley, E. Stride, M.K. Kuimova, Proceed. Nat. Acad. Sci. USA, 110 (2013) 9225-9230. [25] A. Kovalenko, P. Polavarapu, J.-L. Gallani, G. Pourroy, G. Waton, M.P. Krafft, ChemPhysChem, 15 (2014) 2440-2444. [26] S. Dicker, M. Mleczko, M. Siepmann, N. Wallace, Y. Sunny, C.R. Bawiec, G. Schmitz, P. Lewin, S.P. Wrenn, Ultrasound Med. Biol., 39 (2013) 1292-1302. [27] N. de Jong, A. Bouakaz, P. Frinking, Echocardiography, 19 (2002) 229-240. [28] E. Kimmel, B. Krasovitski, A. Hoogi, D. Razansky, D. Adam, Ultrasound Med. Biol., 33 (2007) 1767-1776. [29] M.R. Sprague, E. Chérin, D.E. Goertz, F.S. Foster, Ultrasound Med. Biol., 36 (2010) 313-324. [30] J. Alter, C.A. Sennoga, D.M. Lopes, R.J. Eckersley, D.J. Wells, Ultrasound Med. Biol., 35 (2009) 976-984. [31] S.R. Sirsi, M.A. Borden, Theranostics, 2 (2012) 1208-1222.

Page 28 of 31

26 [32] M.A. Borden, D.E. Kruse, C.F. Caskey, S. Zhao, P.A. Dayton, K.W. Ferrara, IEEE Trans. Ultrasound Ferroelectr. Freq. Control, 52 (2005) 1992-2002. [33] F. Gerber, M.P. Krafft, G. Waton, T.F. Vandamme, New J. Chem., 30 (2006) 524-527. [34] M.P. Krafft, M. Goldmann, Curr. Opin. Colloid Interf. Sci., 8 (2003) 243-250.

ip t

[35] C. Szijjarto, S. Rossi, G. Waton, M.P. Krafft, Langmuir, 28 (2012) 1182-1189. [36] M.J. Landsberg, J.L. Ruggles, W.M. Hussein, R.P. McGeary, I.R. Gentle, B. Hankamer, Langmuir, 26 (2010) 18868-18873

cr

[37] P. Scheibe, J. Schoenhentz, T. Platen, A. Hoffmann-Röder, R. Zentel, Langmuir, 26 (2010) 18246-18255 E. Górecka, Chem. Commun., 46 (2010) 1896-1898. [39] M.P. Krafft, Acc. Chem. Res., 45 (2012) 514-524.

us

[38] P. Nitoń, A. Żywociński, R. Holyst, R. Kieffer, C. Tschierske, J. Paczesny, D. Pociecha,

an

[40] R.C. Buck, J. Franklin, U. Berger, J.M. Conder, I.T. Cousins, P. de Voogt, A.A. Jensen, K. Kannan, S.A. Mabury, S.P.J.V. Leeuwen, Integr. Environ. Assess. Manag., 7 (2011) 513–541.

M

[41] Z. Wang, I.T. Cousins, M. Scheringer, R.C. Buck, K. Hungerbühler, Environ. Int., 70 (2014) 62-75.

d

[42] OECD, Organisation for Economic Co-operation and Development, 2014. OECD/UNEP Global PFC Group, Synthesis paper on per- and polyfluorinated chemicals

te

(PFCs), Environment, Health and Safety, Environment Directorate, OECD. http://www.oecd.org/env/ehs/risk-management/synthesis-paper-on-per-and-polyfluorinated-

Ac ce p

chemicals.htm

[43] M.P. Krafft, J.G. Riess, Chemosphere, doi: 10.1016/j.chemosphere.2014.08.039 (2014).

[44] J.G. Riess, Chem. Rev., 101 (2001) 2797-2920. [45] F. Kiessling, S. Fokong, P. Koczera, W. Lederle, T. Lammers, J. Nucl. Med., 53 (2012) 345-348.

[46] G. Pu, M.A. Borden, M.L. Longo, Langmuir, 22 (2006) 2993-2999. [47] S. Kvåle, H.A. Jacobsen, O.A. Asbjørnsen, T. Omtveit, Sep. Technol., 6 (1996) 219226. [48] M.A. Wheatley, F. Forsberg, N. Dube, M. Patel, B.E. Oeffinger, Ultrasound Med. Biol., 32 (2006) 83-93. [49] J.A. Feshitan, C.C. Chen, J.J. Kwan, M.A. Borden, J. Colloid Interf. Sci., 329 (2009) 316-324. [50] K.P. Pancholi, U. Farook, R. Moaleji, E. Stride, M.J. Edirisinghe, Eur. Biophys. J., 37 (2008) 515-520.

Page 29 of 31

27 [51] E. Stride, M.J. Edirisinghe, Soft Matter, 4 (2008) 2350-2359. [52] E. Stride, M.J. Edirisinghe, Med. Biol. Eng. Comput., 47 (2009) 809-811. [53] U. Farook, H.B. Zhang, M. Edirisinghe, E. Stride, N. Saffari, Med. Eng. Phys., 29 (2007) 749-754.

ip t

[54] E. Talu, K. Hettiarachchi, R.L. Powell, A.P. Lee, P.A. Dayton, M.L. Longo, Langmuir, 24 (2008) 1745-1749. [55] http://www.optisonimaging.com.

cr

[56] S. Tinkov, R. Bekeredjian, G. Winter, C. Coester, J. Pharm. Sci., 98 (2009) 1935-1961. [57] S.J. Satinover, J.D. Dove, M.A. Borden, Ultrasound Med. Biol., 40 (2014) 138–147.

us

[58] S. Rossi, C. Szíjjártó, F. Gerber, G. Waton, M.P. Krafft, J. Fluorine Chem., 132 (2011) 1102-1109.

[59] D. Chatterjee, K. Sarkar, P. Jain, N. Schreppler, Ultrasound Med. Biol., 31 (2005) 781-

an

786.

[60] M.-X. Tang, R. Eckersley, J. Noble, Ultrasound Med. Biol., 31 (2005) 377-384. [61] N. de Jong, L. Hoff, T. Skotland, N. Bom, Ultrasonics, 30 (1992) 95-103.

M

[62] P. Frinking, N. de Jong, Ultrasound Med. Biol., 24 (1998) 523-533. [63] D.E. Goertz, N. de Jong, A.F.W. van der Steen, Ultrasound Med. Biol., 33 (2007) 1376-

d

1388.

[64] J. Benjamins, A. Cagna, E.H. Lucassen-Reynders, Colloids Surf. A, 114 (1996) 245.

6404-6408.

te

[65] P.N. Nguyen, G. Waton, T. Vandamme, M.P. Krafft, Angew. Chem. Int. Ed., 52 (2013)

Ac ce p

[66] D. Langevin, Annu. Rev. Fluid Mech., 46 (2014) 47-65. [67] L. Liggieri, R. Miller, Curr. Opin. Colloid Interface Sci., 15 (2010) 256–263 [68] F. Giulieri, F. Jeanneaux, M. Goldmann, M.P. Krafft, Langmuir, 28 (2012) 1202212029.

[69] J.J. Kwan, M.A. Borden, Langmuir, 26 (2010) 6542-6548. [70] F. Gerber, M.P. Krafft, T.F. Vandamme, M. Goldmann, P. Fontaine, Angew. Chem. Int. Ed., 44 (2005) 2749-2752.

[71] G. Pu, M.L. Longo, M.A. Borden, J. Am. Chem. Soc., 127 (2005) 6524-6525. [72] M. Borden, Soft Matter, 5 (2009) 716-720. [73] H. Nakahara, M.P. Krafft, A. Shibata, O. Shibata, Soft Matter, 7 (2011) 7325-7333. [74] A. Kovalenko, Ph.D. Dissertation, University of Strasbourg, (2013). [75] G.L. Gaines, Insoluble Monolayers at Liquid-Gas Interfaces, Interscience, New York, 1966. [76] A.G. Overbeck, D. Möbius, J. Phys. Chem. , 97 (1993) 7999-8004. [77] A.M. Bibo, C.M. Knobler, I.R. Peterson, J. Phys. Chem., 95 (1991) 5591-5599.

Page 30 of 31

28 [78] R.M. Kenn, C. Böhm, A.M. Bibo, I.R. Peterson, H. Möhwald, J. Als-Nielsen, K. Kjaer, J. Phys. Chem., 95 (1991) 2092-2097. [79] M. Maaloum, P. Muller, M.P. Krafft, Angew. Chem. Int. Ed., 41 (2002) 4331-4334. [80] P.N. Nguyen, T.T. Trinh Dang, G. Waton, T. Vandamme, M.P. Krafft, ChemPhysChem,

ip t

12 (2011) 2646-2652. [81] P.N. Nguyen, M. Veschgini, M. Tanaka, G. Waton, T. Vandamme, M.P. Krafft, Chem. Commun., 50 (2014) 11576-11579.

cr

[82] P.N. Nguyen, G. Nikolova, P. Polavarapu, G. Waton, L.T. Phuoc, G. Pourroy, M.P. Krafft, RSC Adv., 3 (2013) 7743-7746.

us

[83] M. Schmutz, B. Michels, P. Marie, M.P. Krafft, Langmuir, 19 (2003) 4889-4894. [84] J.G. Riess, C. Cornélus, R. Follana, M.P. Krafft, A.M. Mahé, M. Postel, L. Zarif, Adv. Exp. Med. Biol., 345 (1994) 227-234.

[86] M.P. Krafft, Biochimie, 94 (2012) 11-25.

an

[85] M.P. Krafft, J.G. Riess, Angew. Chem. Int. Ed. Engl., 33 (1994) 1100-1101.

ChemBioChem, 7 (2006) 1160-1163.

M

[87] M. Sanchez-Dominguez, M.P. Krafft, E. Maillard, S. Siegrist, A. Belcourt,

[88] E. Maillard, M.T. Juszczak, A. Langlois, C. Kleiss, M.C. Sencier, W. Bietiger, M.S.

Ac ce p

te

657-669.

d

Dominguez, M.P. Krafft, P.R.V. Johnson, M. Pinget, S. Sigrist, Cell Transplant., 21 (2011)

Page 31 of 31