d Technical Note

Ultrasound in Med. & Biol., Vol. 41, No. 12, pp. 3253–3258, 2015� 2015 The Authors. Published by Elsevier Inc. on behalf of World Federation for Ultrasound in Medicine & Biology. This is

an open access article under the CC BY license (http://creativecommons.org/licenses/by/4.0/).Printed in the USA

0301-5629http://dx.doi.org/10.1016/j.ultrasmedbio.2015.07.024

TECHNIQUE FOR THE CHARACTERIZATION OF PHOSPHOLIPIDMICROBUBBLES COATINGS BY TRANSMISSION ELECTRON MICROSCOPY

JOSHUA OWEN and ELEANOR STRIDEInstitute of Biomedical Engineering, Department of Engineering Science, University of Oxford, Oxford, UK

(Received 9 January 2015; revised 16 July 2015; in final form 24 July 2015)

Abstract—Gas microbubbles stabilized by a surfactant or polymer coating are of considerable clinical interestbecause of their imaging and drug delivery potential under ultrasound exposure. The utility of microbubbles for agiven application is intrinsically linked to their structure and stability. These in turn are highly sensitive to coatingcomposition and fabrication techniques. Various methods including fluorescence and atomic force microscopy havebeen applied to characterizemicrobubble properties, but direct observation of coating structure at the nanoscale stillposes a considerable challenge. Here we describe a transmission electronmicroscopy (TEM) technique to observe thesurface of microbubbles. Images from a series of phospholipid-coated microbubble systems, including those deco-rated with nanoparticles, are presented. They indicate that the technique enables visualization of the coating struc-ture, in particular lipid discontinuities and nanoparticle distribution. This information can be used to betterunderstand how microbubble surface structure relates to formulation and/or processing technique and ultimatelyto functionality. (E-mail: eleanor.stride@eng.ox.ac.uk) � 2015 The Authors. Published by Elsevier Inc. on behalfof World Federation for Ultrasound in Medicine & Biology. This is an open access article under the CC BY license(http://creativecommons.org/licenses/by/4.0/).

Key Words: Microbubbles, Ultrasound contrast agent, Nanoparticles, Transmission electron microscopy, TEM.

Microbubbles consisting of a gas core surrounded by acoating of a biocompatible surfactant or polymer arecurrently used in ultrasound imaging as contrast agents(Harvey et al. 2002; Meltzer et al. 1980). They have beenreported to move through the vasculature in a mannerkinetically similar to erythrocytes (Jayaweera et al.1994; Keller et al. 1989) and can improve ultrasoundbackscatter from blood by several orders of magnitude(Gramiak and Shah 1968). The outer coating stabilizesthe microbubble by providing a barrier to gas diffusionand reducing surface tension (Nanda 1997). Its structureand mechanical properties have a strong influence on thebubble’s acoustic response and destruction thresholdand hence utility in clinical applications (Stride 2008;Stride and Edirisinghe 2008). It also provides a surfaceenabling therapeutic material to be attached to the bubbleso that it can be used as a vehicle for drug delivery orgene therapy (Lentacker et al. 2007; Unger et al. 1998).

Microbubbles may be functionalized by the additionof other species such as targeting ligands or nanoparticles

Address correspondence to: Dr. Eleanor Stride, Institute ofBiomedical Engineering, University of Oxford, Old Road CampusResearch Building, Oxford OX3 7 DQ, UK. E-mail: eleanor.stride@eng.ox.ac.uk

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to enable localization and/or multi-modal imaging, suchas via magnetic resonance and photoacoustics (Doveet al. 2013; Yang et al. 2009). In order to optimize theircharacteristics for a given application, it is important tobe able to determine the quantity of a functional specieson a bubble and their location and distribution (Ferraraet al. 2009). Hence, there is a significant need for directvisualization of the nanoscale structure of the microbub-ble shell.

The most widely used method for examiningmicrobubble surface structure, such as the distributionof surface ligands, is via fluorescence microscopy (e.g.,Borden et al. 2006). However, as highlighted byKooiman et al. (2014), this is only suitable for relativelylarge bubbles, outside the clinically relevant range. Fluo-rescence lifetime imaging enables quantification ofnano-scale surface properties but not direct observationof spatial features (Hosny et al. 2013). Polymer-coatedmicrobubbles can be readily visualized via electronmicroscopy because of their relatively stiff shell (Heet al. 2012; Yang et al. 2009). Similarly, liposomalstructures have been widely analyzed via electronmicroscopy (Amstad et al. 2011; Voinea et al. 2005),and these techniques have been adapted to imageacoustically active liposomes (i.e., bi- or multi-lamellar

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phospholipid coated particles that contain gas pockets).May et al. obtained scanning electron micrographs ofacoustically active liposomes (diameter . 1 mm) frozenand cracked via a cryostage and successfully measuredthe shell thickness (May et al. 2002). Vlaskou et al.embedded magnetic and acoustically active liposomes(diameter . 1 mm) in 10% gelatin, and thin sectionswere then stained with uranyl acetate and lead citratefor contrast to study the distribution of nanoparticleswithin the liposome structure (Vlaskou et al. 2010).Hitchcock et al. examined echogenic liposomes 400nm in diameter also using negative uranyl acetate stain-ing using sufficiently high magnification that the phos-pholipid bilayer was visible. Voids corresponding togas pockets were successfully observed, indicating thatthe liposomes remained structurally intact in the vacuumenvironment of the electron microscope (Hitchcock et al.2010).

Phospholipid coated microbubbles (i.e., microscalegas spheres surrounded by a phospholipid monolayer)are more challenging to image, however, because oftheir tendency to collapse under vacuum. Abciximabimmunobubbles developed by Bracco Research SA(Geneva, Switzerland) were successfully observed viascanning electron microscopy (SEM) bound to athrombus after drying and coating with a gold/palladiummixture (Metzger et al. 2015). This technique is, howev-er, specific for bound microbubbles, and nano-scalesurface details could not be observed in the images.Transmission electron microscopy (TEM) was alsoused to confirm attachment of a gold labeledantibody to the surface of a Visualsonics Micromarker(FUJIFILM Visualsonics Inc., Toronto, Ontario, Can-ada) target ready contrast agent (Martin et al. 2007).However, it was not stated whether or not microbubbleswere stabilized in the vacuum environment and againnanoscale surface structures could not be clearlydiscerned. Kim et al. used freeze-fracture TEM toobtain images of a replica of a phospholipid-coatedmicrobubble surface, which showed ‘‘domain-like’’structures with sub-micrometer dimensions (Kim et al.2003). This technique was also utilized by Bordenet al. and Sirsi et al. to observe surface structures onphospholipid-coated microbubbles and lung surfactantmicrobubbles, respectively (Borden et al. 2006; Sirsiet al. 2009). However, freeze-fracture TEM is an expen-sive technique and has not been commonly used formicrobubble characterization.

The aim of this study was to determine whetherphospholipid-coated microbubbles could be imagedintact using conventional TEM by modifying the samplepreparation method. We report a simple, low-cost tech-nique to concentrate the microbubbles on the electronmicroscope grid by exploiting their buoyancy followed

by uranyl acetate staining both to provide contrast andto assist in maintaining the structural integrity of themicrobubble.

METHODS

Three types of phospholipid microbubble withdifferent surface characteristics were analyzed: (i)microbubbles coated with a mixture of 1,2-Distearoyl-sn-glycero-3-phosphocholine (DSPC) and polyethylenegly-col-40-stearate (9:1 molar ratio); (ii) microbubbles coatedwith the same phospholipid/emulsifier mixture combinedwith 15nmspherical gold nanoparticles (23 1014 nanopar-ticles/mL) (Mohamedi et al. 2012); and (iii) phospholipidmicrobubbles mixed with liquid droplets containing 10nm magnetic nanoparticles (5 3 1013 nanoparticles/mL)(Owen et al. 2012; Stride et al. 2009). All microbubbleswere prepared by sonication in deionized water usingsulphur hexafluoride (SF6) as the filling gas. Please seereferences for further fabrication details. DSPC waspurchased from Avanti Polar Lipids (Alabaster, Alabama,USA), magnetic nanoparticles from Liquids Research(Bangor, UK), and all other materials from Sigma-Aldrich (Dorset, UK).

For transmission electron microscopy, 30 min aftermicrobubble production, 5 mL of each microbubble solu-tion (�109 bubbles/mL) was applied to carbon film–coated 300 mesh copper grids (Electron MicroscopySciences, Hatfield, PA, USA), which had been gentlyionized in a plasma cleaner (Harrick Plasma, Ithaca,NY) for 30 s. The first grid was used as a control, andphospholipid microbubbles were simply added in asimilar manner to liposomes (Hitchcock et al. 2010),whereas the second grid was inverted for approximately1 min to allow a high concentration of microbubbles toaccumulate on the grid surface. This process was repeatedfor all microbubble samples. Grids were negativelystained by incubation with a 5 mL drop of 2% w/v uranylacetate for 30 s. The grid was then dried using filter paperand left for approximately 1 min. The grids could then bestored indefinitely for analysis. Samples were visualizedat 80 kV with an FEI Tecnai T12 electron microscopeand low-dose images were acquired at �0.8 mm underfo-cus with 15 e2/�A2 on an FEI Eagle CCD camera. Imageswere acquired at a nominal magnification of 346000,which was found to be at a pre-calibrated magnificationof 356603 to produce a specimen sampling of 0.265nm/pixel.

RESULTS

Phospholipid microbubbles (9:1 DSPC:PEG-40-Stearate)

No microbubbles could be discerned on the controlgrid (non-inverted sample). Instead objects of various

Fig. 2. Transmission electron microscope image of a pre-concentrated sample of phospholipid-coated microbubbles. Ar-row 1 indicates a large discontinuity on themicrobubble surface.The inset shows a higher magnification image of the microbub-ble and arrow 2 indicates a lipid coated object on the surface ofthe microbubble. The dark areas surrounding the bubbles are

due to pooling of the uranyl acetate stain.

TEM characterization of phospholipid microbubble coatings d J. OWEN and E. STRIDE 3255

shapes and sizes were seen (Fig. 1). Previous electronmicroscopy observations of liposomes (Hitchcock et al.2010) via a similar method suggest that these werenon-buoyant lipid-coated objects generated during themicrobubble production process. When microbubbleswere concentrated on the copper grid by inversion, how-ever, the TEM images clearly showed circular objectsbetween 1 and 5 mm in diameter (Fig. 2), consistentwith the microbubble size distribution obtained from op-tical microscopy (supplementary data). Increasing themagnification revealed large discontinuities in the phos-pholipid shell (Fig. 2, arrow 1), which may correspond tofolds in the shell, and also the presence of discrete struc-tures of similar contrast, possibly vesicles, preferentiallyattached to the surface near these discontinuities (Fig. 2,arrow 2).

Phospholipid microbubbles coated with goldnanoparticles

Again the control sample had no intact microbub-bles, whereas the inverted grid revealed microbubblesthat exhibited fewer discontinuities across their surfacesrelative to the DSPC:PEG-40-stearate microbubbles. Athigher magnification, high-contrast objects wereobserved on the surface consistent in size and contrastwith the gold nanoparticles (Fig. 3, arrow 1). Those dis-continuities that could be observed on the surface weremuch less pronounced than on DSPC:PEG-40-stearatemicrobubbles (arrow 2). Slightly larger, lower contrastcircular regions (arrow 3) could also be observed in

Fig. 1. Transmission electron microscope image of a sample ofphospholipid coated microbubbles where the grid was not

inverted.

Fig. 3. Transmission electron microscope image of a pre-concentrated sample of phospholipid microbubbles coated ingold. The inset shows a higher magnification image of the mi-crobubble surface. Arrow 1 indicates a gold nanoparticle 10nm in diameter. Arrow 2 indicates a discontinuity in the phos-pholipid surface and arrow 3 is an example of a higher contrastcircular region, which are seen over the whole microbubble

surface.

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high concentration over the whole microbubble surface.These could be domains of condensed or expanded lipidformed during microbubble cooling (analogous to grainboundaries in crystalline materials [Kim et al. 2003])or, more likely, gold nanoparticles displaced from thefocal plane. Analysis of the images (8 separate microbub-bles) indicated that the gold nanoparticles were randomlydistributed over the microbubble surface with a surfaceconcentration of approximately 13 6 2 nanoparticles/mm2

(taking into account only those particles that were infocus in the images). This is consistent with anadsorption-based attachment mechanism. From the con-centration and size distribution of the microbubbles(supplementary data), this corresponds to 5 3 108 nano-particles per mL of the suspending solution becomingbound to the microbubbles (i.e., ,1% of the availableparticles [2 3 1014 nanoparticles/mL]).

Magnetic droplets and phospholipid microbubblesFigure 4 shows DSPC:PEG-40-stearate (9:1) micro-

bubbles mixed with magnetic droplets examined via elec-tron microscopy. Again discontinuities in the coatingcould be seen and, at higher magnification, clusters ofiron oxide nanoparticles were visible at these discontinu-ities (Fig. 4, inset) consistent with adsorption of magneticdroplets onto the surface of the microbubbles. The size ofdroplets varied, as did the number of droplets attached toeach microbubble, but on average (based on 8 separate

Fig. 4. Transmission electron microscope image of a pre-concentrated sample of phospholipid microbubbles mixedwith magnetic droplets. The inset shows a higher magnificationimage of the microbubble surface with clusters of nanoparticles

located at a surface discontinuity.

bubbles) therewere 3 droplets per 10 mm2with�20 nano-particles each. This again corresponded to a small propor-tion of the nanoparticles available in the suspendingliquid (,1%).

DISCUSSION

In the inverted samples, objects with a circularprojection consistent with intact microbubbles wereobserved from all the formulations. The sizes of themicrobubbles observed via electron microscopy corre-sponded to those obtained via optical microscopy(supplementary data). This was in contrast to the controlsamples in which no regularly shaped objects were seen(Fig. 1). Nano-scale features of the microbubble surfacewere clearly visible in all cases. In those formulationscontaining nanoparticles, dark spherical objects in thecorrect size range could be clearly visualized and theirconcentration and surface distribution could be esti-mated. The technique thus has the potential to test the val-idity of hypothesized bubble structures. For example,solid nanoparticles adsorbed onto the surface of micro-bubbles have been found to influence both their stabilityand acoustic properties (Stride 2008). It has been pro-posed that this is due to a high surface concentrationof particles ‘‘jamming’’ and preventing the bubble fromcompressing, but no direct evidence has previouslybeen obtained. Figure 3 confirms that gold nanoparticlesare indeed located on the surface of the microbubble thatwould enable particle jamming. Similarly, previous workhas indicated that magnetic microbubbles can be formedvia fusion of droplets containing magnetic nanoparticleswith phospholipid-coated microbubbles. Figure 4 pro-vides further evidence for this process, reinforcing resultsobtained previously using fluorescence microscopy andmagnetic targeting (Owen et al. 2012). The results thusindicate that the technique was successful in providinga means of imaging microbubble coating structure atthe nano scale.

LimitationsAs presented, this technique only provides a 2-D

projection of a 3-D structure, which inevitably limitswhat can be observed and deduced. However, the samplepreparation method should be compatible with 3-D imag-ing techniques in a different TEM system. The resultsobtained indicate that microbubbles survive the process-ing technique, but it is unknown whether there are anyother effects on the structure. For example, all microbub-bles had discontinuities within the coating. The phospho-lipid microbubbles had what appeared to be folds in theirshell. These could have been due to some degree of crum-pling in the electron microscope vacuum or simplymicro-bubbles losing gas and wrinkling as they shrank before

TEM characterization of phospholipid microbubble coatings d J. OWEN and E. STRIDE 3257

imaging. Alternatively they may have been related to thedomain boundaries reported by Kim et al. (2003).

Objects in the solution that are reversibly bound tothe microbubble surface cannot be differentiated fromobjects that are irreversibly bound, such as vesicles ornanoparticles. Centrifugation could be used in order toremove any non-bound objects before imaging to addressthis question (Feshitan et al. 2009), although it is likelysome will still be present in solution. Nanoparticle con-centration and surface distribution can be deduced fromthe images; at present, however, there is no other tech-nique available to confirm the results obtained.

CONCLUSIONS

A simple technique has been developed to enablevisualization of gas-filled microbubbles within the vac-uum environment of an electron microscope. Exploitingthe microbubbles’ buoyancy to achieve a high concentra-tion on the sample grid and coating the microbubbles withuranyl acetate allows their structure to be maintainedduring imaging. The technique allows surface featuressuch as discontinuities and vesicles to be observed onphospholipid microbubbles. In addition, functional com-ponents such as nanoparticles embedded within the bub-ble shell can be directly visualized. The technique is lowcost and versatile and it is hoped will be a useful tool formicrobubble characterization, for example, to examinethe relationship between shell structure, compositionand fabrication technique.

Acknowledgments—We would like to record our sincerest thanks toDr. Alistair Siebert and Prof. Kay Gr€unewald for their expert adviceand technical assistance in developing the technique.

We would like to thank the Engineering and Physical SciencesResearch Council for supporting the work through EP/I021795/1. Sup-porting data are available through the University of Oxford ORA datarepository (doi: http://10.5287/bodleian:xp68kh14d).

SUPPLEMENTARY DATA

Supplementary data related to this article can be found at 10.1016/j.ultrasmedbio.2015.07.024.

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