Published: October 25, 2011

r 2011 American Chemical Society 15024 dx.doi.org/10.1021/la202679p | Langmuir 2011, 27, 15024–15033

ARTICLE

pubs.acs.org/Langmuir

Silica-CoatedQuantumDots for Optical Evaluation of PerfluorocarbonDroplet Interactions with CellsIvan Gorelikov,† Amanda L. Martin,† Minseok Seo,†,‡ and Naomi Matsuura*,†,‡

†Imaging Research, Sunnybrook Research Institute, and ‡Department of Medical Biophysics, University of Toronto, 2075 Bayview Avenue,Toronto, Ontario, Canada M4N 3M5

bS Supporting Information

ABSTRACT:

There has been recent interest in developing new, targeted, perfluorocarbon (PFC) droplet-based contrast agents for medicalimaging (e.g., magnetic resonance imaging, X-ray/computed tomography, and ultrasound imaging). However, due to the largenumber of potential PFCs and droplet stabilization strategies available, it is challenging to determine in advance the PFC dropletformulation that will result in the optimal in vivo behavior and imaging performance required for clinical success. We propose that theintegration of fluorescent quantum dots (QDs) into new PFC droplet agents can help to rapidly screen new PFC-based candidateagents for biological compatibility early in their development. QD labels can allow the interaction of PFC droplets with single cells tobe assessed at high sensitivity and resolution using optical methods in vitro, complementing the deeper depth penetration but lowerresolution provided by PFC droplet imaging using in vivomedical imaging systems. In this work, we introduce a simple and robustmethod tomiscibilize silica-coated nanoparticles into hydrophobic and lipophobic PFCs through fluorination of the silica surface viaa hydrolysis-condensation reaction with 1H,1H,2H,2H-perfluorodecyltriethoxysilane. Using CdSe/ZnS core/shell QDs, we showthat nanoscale, QD-labeled PFC droplets can be easily formed, with similar sizes and surface charges as unlabeled PFC droplets. TheQD label can be used to determine the PFC droplet uptake into cells in vitro by fluorescencemicroscopy and flow cytometry, and canbe used to validate the fate of PFC droplets in vivo in small animals via fluorescence microscopy of histological tissue sections. This isdemonstrated in macrophage and cancer cells, and in rabbits, respectively. This work reveals the potential of using QD labels forrapid, preclinical, optical assessment of different PFC droplet formulations for their future use in patients.

’ INTRODUCTION

Perfluorocarbons (PFCs) are fluorinated aliphatic compoundsthat have a long history in biomedicine.1�3 The unique proper-ties of PFCs, including their low solubility in water, fats, andlipids; chemical and biological stability; and oxygen-carryingcapacity, have led to many decades of nanoscale (∼100�400 nm) PFC droplet development, culminating in their use inthousands of patients as blood substitute agents.4�8 Despitepromising early clinical investigations of PFCs as contrast agentsfor magnetic resonance imaging (MRI),9 radiography/CT,10,11

and ultrasound imaging,10,12 the use of PFC droplets for medicalimaging in patients has been limited, in part due to the largevolumes of PFC droplets required for successful medical imag-ing, and the need for further optimization of these PFC dropletsfor human injection.13,14

It has recently been shown that the contrast agent dosesrequired for medical imaging may be greatly reduced through thecombination of more sensitive medical imaging systems and theuse of nanoscale agents that can preferentially target disease (e.g.,cancer) sites in vivo, for example, via passive accumulation in theleaky vasculature of tumors15,16 or through active targeting totheir biological ligands.17,18 PFC droplet agent optimization anddose reduction may lead to a decrease in the incidence of seriousadverse effects that have been observed in a small number ofpatients during early trials, including thrombocytopenia, com-plement activation and cytokine release, RES blockade, flu-like

Received: March 8, 2011Revised: September 21, 2011

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symptoms, and central nervous system effects.5,8,19 In addition tothe demand for new targeted imaging agents, continuing advancesin cancer biomarker development,20�22 in vivo nanoscale agenttargeting methods,23 and PFC droplet stabilization strategies24,25

have led to renewed interest in developing nanoscale PFC dropletsfor medical imaging,26�28 presenting materials scientists withexciting opportunities to re-engineer PFC droplet formulationsfor future targeted clinical contrast imaging applications.

However, similar issues related to PFC agent development forhuman use as encountered decades ago remain:5,29�31 given thenumerous available PFCs and stabilization strategies (e.g.,fluorosurfactants,24,25 polymers,32,33 proteins,34 and lipids,35�37

with andwithout targeting groups35�38), how does one rationallydesign the optimal nanoscale PFC droplet agent with appropriatein vivo circulation times to target disease sites, with idealpharmacokinetics to clear from the patient after imaging, andwith suitable imaging properties for the modality and disease sitein question? Although extensive studies of PFC droplets forhuman injection have been conducted, these focused mainly onthe use of PFC droplets as blood substitute agents in whichoxygen solubility was a primary criterion,5,31 permitting the useof non-imaging-specific, high molecular weight/boiling point,PFC-based formulations (i.e., with bp ranging from ∼128 to243 �C).5 New stabilization strategies are required for relativelyunstable, low molecular weight/boiling point PFC formulations39

needed for certain imaging applications (e.g., phase-shift emul-sions for ultrasound imaging and therapy32,40,41) and desirable ingeneral for their more favorable clearance characteristics in vivofrom tissue.39 Since the combination of PFC type and emulsifierdetermines thedroplet size, stability, charge, in vivobiodistribution,31,42

and targeting ability,35�38 ultimately each new PFC droplet formula-tion must be tested individually to assess their modality-dependentimaging performance and targeting potential.

Rapid techniques to screen and select promising PFC dropletformulations from the large number of candidate formulationsfor potential clinical translation are required. Initial assessmentsof biological suitability can be simply and effectively conductedby evaluating a panel of candidate PFC droplets with cellsin vitro.42 After in vitro screening, PFC droplets can be assessedin vivo in small animals using relatively inexpensive, preclinicalimaging systems to evaluate PFC droplet targeting, biodistribu-tion, pharmacokinetics, and imaging contrast. Although standardpreclinical imaging systems (e.g., MRI, CT, or ultrasound) canprovide high depth penetration for in vivo evaluation, they haveinadequate spatial resolution (.0.1 mm) and sensitivity toimage nanoscale PFC droplets interacting with individual cells.43

To complement the lower spatial resolution and sensitivity buthigh depth penetration of in vivo medical imaging, opticalmicroscopy, with high spatial resolution (∼200 nm) and highsensitivity that permits imaging of nanoscale fluorescent agentsin single cells in vitro but at shallow depths (∼0.1 mm), can beused. To facilitate optical microscopy of nonfluorescent PFCdroplets, we propose that labeling the PFC droplet core withhighly fluorescent, nonphotobleaching quantum dot (QD) labels(Figure 1) can provide sensitive, high-resolution optical imagingof PFC droplets during two critical stages of the PFC dropletdevelopment process. At an early stage, the QD label can helpscreen a panel of candidate PFC formulations in relation to targetand nontarget cells in vitro. Simple tests evaluating how rapidlyPFC droplets are phagocytosed by macrophages can help predictin vivo circulation times for passive targeting, while differentactive targeting strategies of PFC droplets can be evaluated

in vitro using target cells.16 At a later stage in preclinicaldevelopment using animal models, the QD label can validatetargeting of the PFC droplet to disease sites and determinebiodistribution in nontarget tissues via optical microscopy ofexcised/biopsied animal tissue.

In this work, we show that QD integration into PFC dropletscan be used to optically evaluate the interactions of PFC dropletswith target cells in vitro and in tissue sections after in vivoinjection, at very high sensitivity and resolution to complementthe in vivo strengths of clinical imaging. Specifically, we introducea simple method of overcoming QD immiscibility within hydro-phobic and lipophobic PFCs by coating the QDs with a thin layerof silica followed by fluorination. After QD miscibilization intoPFCs, we demonstrate that the QD-PFC solution can beemulsified to form different QD-incorporated PFC droplets,without affecting the size or surface charge of the PFC droplets.The QD label can be used to characterize the uptake of differenttypes of PFC droplets in cells to assess their suitability fortargeting disease sites. We further show that the QD labels canbe used to assess PFC droplet distribution in preclinical animalmodels, by sensitive detection of the QDs in histological sectionsof unstained target and nontarget tissues.

’EXPERIMENTAL SECTION

Materials. PFCs (i.e., FC43 (perfluorotributylamines, (CF3(CF2)3)3N,bp 174 �C at 1 atm), FC72 (perfluorohexanes, C6F14, bp 56 �C at 1 atm),perfluoropentane (C5F12, bp 29 �C at 1 atm), FC84 (perfluoroheptanes,C7F16, bp 80 �C at 1 atm), and PFOB (perfluorooctylbromide, C8F17Br, bp142 �C at 1 atm)) were purchased from Synquest Laboratories. ZonylFSP and FSO fluorosurfactants were purchased from Sigma-Aldrich. Allchemicals for silica-coated QD and silica-coated Au NR synthesis and PFCmiscibilization (octadecene, cadmium oxide, selenium, stearic acid, tributyl-phosphine, trioctylphosphine oxide, octadecylamine, cetyltrimethyl ammo-nium bromide, sodium borohydride, NH4OH, tetraethyl orthosilicate,ascorbic acid, silver nitrate, tetrachloroauric acid, and 1H,1H,2H,2H-perfluorodecyltriethoxysilane) were purchased from Sigma-Aldrich,and used as-received. Deionized water (Millipore Milli-Q grade, 18.2 MΩ)was used in all experiments.Methods. Silica-Coating and Miscibilization of Nanoparticles (NPs)

into PFCs. CdSe/ZnS core/shell QDs, capped with TOPO and octadecyl-amine, were synthesized according to a previous method.44 Smooth silicacoating of QDs was achieved using reverse microemulsion.45 Briefly,QDs were dispersed in 100 mL of cyclohexane at a concentration of0.1μM, followed by the addition of 4.6 g of Igepal CO-520. Under vigorousstirring, 1 mL of concentrated ammonia solution was added. After∼1 min,the stirring rate was decreased and 0.8mLofTEOSwas added. After∼24 h,the silica coating procedure was complete.

Au NRs capped with cetyltrimethyl ammonium bromide (CTAB)were synthesized according to a previously published procedure.46

Figure 1. Integrated QD labels can guide the development of new PFCdroplet formulations for medical imaging by facilitating high spatialresolution (∼200 nm) microscopy of PFC droplets with target cells invitro, to complement the in vivo, but lower resolution (.0.1 mm)assessment of PFC droplets for medical imaging (e.g., MRI, CT/X-ray,and ultrasound).

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Mesoporous silica coating was achieved via direct deposition.47 Briefly,excess CTAB surfactant was first removed from the as-synthesized AuNRs via centrifugation of 15 mL aliquots (10 000 rpm for 30 min). Afterdiscarding the supernatant, the precipitate was dispersed in 10 mL ofMilli-Q water, and 100 μL of 0.1 M NaOH solution was added uponstirring. Following this step, three 30 μL injections of 20% TEOS inmethanol were added under gentle stirring at 30 min intervals, and themixture left to react for 2 days.

For NP miscibilization into PFCs, the surfactant was first removedfrom the as-synthesized silica-coated NPs.47 NPs were first precipitatedout of cyclohexane through the addition of a small amount of methanol(5�7% by volume), followed by centrifugation (Eppendorf 5430 Cen-trifuge). The NPs were redispersed in approximately 25 mL of methanolvia sonication, and washed (i.e., centrifugation at 10 000 rpm for 25 minfollowed by redispersion with methanol) 5 more times. To fluorinate thesilica-coated NPs, 1H,1H,2H,2H-perfluorodecyltriethoxysilane wasadded to concentrated (∼3 μM48), silica-coated NPs in methanol in a1:10 000 ratio (i.e., 120 μL in 7 mL of the QD solution) and stirred for 5min, followed by the addition of 30% ammonia�water solution to a finalconcentration of 0.12 M. After stirring for ∼24 h, the fluorinated silica-coated NPs precipitated out of solution. After the removal of methanol,the surface modified silica-coated NPs were dispersed in PFC bysonication for several minutes, which resulted in a clear solution. TheNP-PFC solution was filtered using a 0.45 μm syringe filter.NP and PFC Droplet Characterization. Fluorescence spectra were

obtained using a Horiba Jobin Yvon FluoroMax-4 Spectrofluorom-eter (Edison, NJ). The hydrodynamic diameters and zeta potentialsof PFC droplets were obtained using a Malvern Zetasizer Nano-ZS3000HS (Worcestershire, UK) instrument. Electron microscopyimages were acquired on a Hitachi S-5200 scanning electron micro-scope (SEM).Preparation of QD-Incorporated PFC in PBS Emulsions. To prepare

nanoemulsions, fluorosurfactant (FSP or FSO, 50 μL) was dissolved in10 mL of DI water by brief sonication in a sonicating bath. To thismixture, QD-PFC (300 μL) was added. The solution was thensuspended in a cool water bath and sonicated with a Branson DigitalS450D Sonifier for 7min of exposure at 10% amplitude (6W), using a 1 son, 1 s off pulse sequence. After sonication, the emulsion was then slowlyfiltered into centrifuge tubes using sterile 0.80 μm polyethersulfonemembrane filters (Pall Life Sciences) to remove any large droplets in thepopulation while minimizing droplet vaporization. Samples were thencentrifuged using an Eppendorf 5430 R Centrifuge at 4000 rpm, 4 �C,for 20 min. The supernatant was removed and the remaining pellet wasredispersed with gentle shaking and brief sonication in sterile phosphatebuffered saline (PBS, Wisent), diluted to 10 mL. For micrometer-scale,QD-incorporated, PFC droplet formation, microfluidics was used.49,50 Athree-inlet microfluidic device (MFD) was used to generate QD-incorporated PFC droplets of different sizes with narrow size distribu-tion. The dispersed phase (i.e., ∼10 μM QDs NPs in PFC) wasintroduced in the central channel, while the continuous phase (i.e., 0.5vol % of FSP or FSO in 40 to 60 wt % of glycerol and DI water mixture)was supplied to the two side channels of the MFD. The flow rate of thedispersed and continuous phase was varied from 0.01 to 0.5 mL/h andfrom 0.1 to 5.0 mL/h, respectively. The sizes of the micrometer-scale,QD-incorporated, PFC droplets were analyzed using Image-Pro Plus(Media Cybernetics, USA) software.Evaluation of Cell Loading with QD-Incorporated PFC Droplets.

Murine alveolar macrophage cells (RAW264.7) and human breast carci-noma cells (MDA-MB231) were received from ATCC (Rockville, MD).RAW264.7 cells were maintained at 37 �C in a humidified atmosphere with5% CO2 in delta-minimum essential medium (DMEM, 4.5 g/L glucose,L-glutamine, and sodium pyruvate, Wisent) supplemented with 10% heat-inactivated fetal bovine serum (FBS, Wisent), while MDA-MB231 cellsweremaintained in RPMI 1640medium (Wisent) supplemented with 10%

FBS. Cells were seeded at 8 � 105 cells/well in 6-well plates andallowed to adhere and grow until 80% confluency was reached. Themedium was aspirated and replaced with 1 mL of fresh medium, towhich 70 μL of the desired emulsion was added. (The emulsion wasadded in excess to reduce the dependence of the uptake of QD-PFCdroplets on the concentration of droplets in the media.) Before QD-PFC droplets were incubated with cells, the fluorescence of eachemulsion was measured, to normalize the fluorescence intensity foreach experiment. The cells were incubated for the desired time,followed by washing with PBS twice to remove excess emulsion.RAW264.7 cells were detached from the plate using a cell scraper(Sarstedt), while MDA-MB231 cells detached using trypsin, followedby centrifugation and redispersion in PBS (0.5 mL). Cells weretransferred into round-bottomed tubes (BD Falcon) containing0.5 mL of 4% paraformaldehyde (PFA) where they were left undis-turbed for 30 min at room temperature. Tubes were centrifuged at3600 rpm at 4 �C for 5 min. The supernatant was removed, and thecells were redispersed in 1 mL of PBS with repeated pipetting. Theamount of PFC labeling was determined by flow cytometry on a BDFACSCalibur flow cytometer (BD Biosciences, USA). Basal fluores-cence was given by cells that were not incubated with the emulsion.Flow cytometry data were gated to exclude small cell debris andanalyzed using FloJo software. Samples were analyzed using filtersoptimized for the emission spectra of the QDs, with the same voltagesetting for each data set. Use of the same voltage setting validatescomparison among different batches, samples, and cell lines. Forwardand side scatter were determined based on the cell line used.

Fluorescence Microscopy of QD-Incorporated PFC Droplets. Cellswere plated on coverslips (BD Biosciences) in 6-well plates andincubated with emulsions as above. The cells were then washed twicewith PBS, and stained in the cold with Alexa488-Concanavalin A(Molecular Probes) to label the cell membrane, then fixed in 4%paraformaldehyde, then stained with DAPI (Sigma) to label nuclei,and mounted in DaKo fluorescent mounting medium (DaKo Cy-tomation). Images were obtained using inverted epifluoresence micro-scopy using a Zeiss Axiovert 200 M microscope with Axiovision imagingsoftware and standard FITC, cy3, and DAPI filter sets.

Transmission Electron Microscopy (TEM) and X-ray Spectroscopy.Cells were plated in 10 mL cell culture dishes and grown to 80%confluency. The desired emulsion (350 μL) was added and incubatedwith the cells for two hours, at which point the medium was aspiratedand the cells washed twice with PBS. Cells were detached using a cellscraper and transferred to a 1.5 mL eppendorf centrifuge tube. Cellswere centrifuged at 3600 rpm at 4 �C for 5 min. The supernatant wasremoved and cells were fixed in 2% glutaraldehyde in 0.1 M sodiumcacodylate buffer, postfixed in 1% osmium tetroxide in the same buffer,dehydrated in a graded ethanol series followed by propylene oxide, andembedded in Quetol-Spurr resin. Sections 100 nm thick were cut on anRMC MT6000 ultramicrotome and viewed in an FEI Tecnai 20 TEMequipped with a Gatan Dualview digital camera. Energy-dispersive X-rayspectroscopy was performed with an EDAX Phoenix system.

In Vivo Demonstration. New Zealand white rabbits (Charles RiverLaboratory) were anesthetized via intramuscular injection using keta-mine and xylazine. 1.5 mL/kg of the nanoscale, FSP-stabilized, QD-C6F14 droplet agent was administered to the anesthetized rabbit at13.5 mL/h into the cannulated ear vein. Twenty-four hours afterinjection, the rabbit was sacrificed and organs were surgically removedand preserved in 40% formalin. The organs were then serially sectionedinto 10-μm-thick slices and alternate sections were stained with hema-toxylin and eosin (H & E), with the remaining sections unstained. Thetissue sections were then examined under bright-field and fluorescencemicroscopy. The animal experiments were conducted in accordancewith the recommendations of the Animal Care Committee, SunnybrookHealth Sciences Centre.

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’RESULTS AND DISCUSSION

Integration and Characterization of Silica-Coated QDsinto PFC Droplets. PFCs are extremely hydrophobic andlipophobic and are immiscible with aqueous and commonorganic solvents.51 Thus, nanoparticles (NPs) as-synthesizedvia standard wet chemistry methods are not miscible in PFCs.We demonstrate a simple and robust method that overcomesQD immiscibility with PFCs by coating the QDs with a thinlayer of silica followed by fluorination. Silica is an extensivelystudied biomaterial that can be easily deposited on a varietyof colloidal NPs synthesized in both organic or aqueousmedia.47,52�60 A previous study has shown that hydrophobic,tri-n-octylphosphineoxide (TOPO)-capped QDs can be incor-porated into PFCs using ligand exchange with 1H,1H,2H,2H-perfluorooctanethiol.61,62 We introduce an alternative, rapid,room-temperature procedure for NP miscibilization into PFCsthat is not limited to QDs or NPs synthesized in organic mediawith thiol-binding surfaces (e.g., containing Zn or Cd)61,62 andnot impacted by thiol-based oxidation that can reduce QDquantum yield and colloidal stability.63 Since the thin silicacoating serves as a common platform for fluorination, there isno limitation on the initial NP synthesis media, and the samemiscibilization procedure is possible using different QDs andNPs for integration into PFC droplets.In this study, CdSe/ZnS QDs were selected for their light

emission in the red visible range, suitable for reducing interferencefrom surrounding tissue autofluorescence and for their compat-ibility with standard filters used in cellular biology (e.g., using flowcytometry and fluorescence microscopy). Hydrophobic,

fluorescent, CdSe/ZnS core/shell QD NPs (∼5 nm in diameter)capped with octadecylamine44 and coated with ∼10-nm-thicksilica layer using reverse microemulsion45 were synthesized(Figure 2a,b). The miscibilization of silica-coated QDs in PFCwas accomplished using a straightforward procedure, in which1H,1H,2H,2H-perfluorodecyltriethoxysilane was reacted with thesilica surface via a hydrolysis-condensation reaction64 (Figure 2c).Fluorination of silica-coated NPs is a simple, single-step processwhere the silane group of perfluorodecyltriethoxysilane reacts withthe silica surface.60 Ammonia was added to increase pH in order tocatalyze the hydrolysis-condensation reaction.65 After this fluor-ination procedure, all the silica-coated NPs precipitated out of theaqueous solution. No measurable QD fluorescence was observedin the water phase, indicating a high (i.e., close to 100%) transferefficiency of silica-coatedQDs to PFC. After fluorination, the QDsare completely immiscible in aqueous or standard organic solvents,while miscible in fluorinated solvents. The silica-coated QDswere miscible in every fluorinated aliphatic compound tested,including perfluorotributylamine (CF3(CF2)3)3N, perfluoropen-tane (C5F12), perfluorohexanes (C6F14), perfluoroheptanes(C7F16), and perfluoroocytlbromide (C8F17Br). The peak fluor-escence of silica-coated QDs did not shift significantly uponredispersion from toluene to water (∼5 nm red-shift), nor uponfluorination and dispersion and redispersion into PFCs (∼5 nmblue-shift) (Supporting Information, Figure S1). The number ofsilica-coatedQDs dispersed in PFCsmay be controlled by addingor evaporating PFCs after the QDmiscibilization procedure. Forhigh PFC droplet labeling and to prevent QD aggregation at highQD concentrations, the QDs were typically dispersed into PFCsto a concentration of ∼10 μM NP.48

The miscibility of the QDs in PFCs before and after fluorina-tion can be clearly observed in Figure 2d. Uniform dispersion ofthe fluorescent, silica-coated QDs in water, prior to surfacefluorination of the silica, phase-separated from the denser PFC(here, C6F14, F = 1.76 g/mL), is shown in the left-hand vial. Aftersurface modification, the QDs become dispersed in PFCs, asshown by themiddle vial in Figure 2d, and completely immisciblein water. Once fluorinated, QDs remained uniformly dispersedand exhibited no observable precipitation within the PFC afterseveral months. This stability is expected due to the fact thatfluorosilanes form up to three stable Si�O�Si bonds during thehydrolysis-condensation reaction.66 After miscibilization of theQDs into PFCs, the QD�PFC solution can be emulsified toform PFC droplets. For example, the addition of an anionicfluorosurfactant into the water and QD-incorporated PFCmixture followed by sonication resulted in the formation ofnanoscale QD�PFC droplets suspended in water, as shown bythe right-hand vial in Figure 2d.The QDs should be loaded inside the droplet, such that the

PFC droplet surface can be optimized for particular biologicalapplications16,67 and/or for biological conjugation to cellulartargets.35�38 Furthermore, internal labeling of the PFC dropletpermits retention of the desired properties after the removal ofpotentially toxic QDs68 prior to human use, after their optimi-zation in animal models. To image the QD distribution withinindividual PFC droplets using optical microscopy, composi-tionally identical, monodisperse, micrometer-scale dropletswere synthesized using the QD�PFC solution and the anionicfluorosurfactant used in Figure 2. No measurable fluorescencearising from free QDs was observed in the continuous phaseoutside the droplets, and all droplets seen in bright-fieldmicroscopy were labeled by QDs (Supporting Information,

Figure 2. Silica-coated CdSe/ZnS QD NPs, ∼5-nm-diameter, coatedwith a ∼10-nm-thick, smooth silica layer: (a) TEM image; (b) SEMimage; (c) schematic of fluorinated silica-coated QDs; and (d) silica-coated QDs photographed under illumination by 365 nmUV light: QDsbefore fluorination dispersed in water (W) and phase-separated from thedenser C6F14 (PFC) (left); QDs after fluorination dispersed in C6F14phase-separated from water (middle); and after formation of QD-labeled C6F14 droplets in water (right).

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Figure S2). Although PFC droplet growth can occur over time,the QDs were never observed to extract from the droplets insaline or PBS, even several weeks after synthesis. The immis-cibility of fluorinated, silica-coated QDs in water and the lack offree QDs observed in water suggests that the concentration ofQDs in the droplets is similar to its concentration in the originalPFC solution (∼10 μM).Ideally, the PFC droplets’ properties should depend only on

the emulsification conditions (i.e., type and concentration ofPFC and emulsifier, as well as the emulsification method), andnot on the NP incorporation into the droplet. To test whetherthe integration of QDs into the PFC affected the size and surfaceproperties of nanoscale PFC droplets, QDs incorporated intotwo different nanoscale PFC droplets (i.e., C6F14, bp 56 �C, and(CF3(CF2)3)3N, bp 170�180 �C) using identical sonicationmethods were synthesized using anionic (Zonyl FSP) andnonionic ethoxylated (Zonyl FSO) fluorosurfactants, and com-pared to PFC droplets not containing QDs. Although any type ofPFC droplet could be selected for these studies, C6F14 and(CF3(CF2)3)3N were selected to demonstrate that this methodof QD inclusion into PFC droplets is applicable to both low-boiling-point and high-boiling-point PFCs. FSP and FSO havesimilar fluorinated tails for PFC droplet stabilization, but theirdifferent headgroups result in different water and PFC solubi-lities and charges. Thus, the use of different fluorosurfactant-PFCcombinations in the same ratios can result in the production ofdroplets with different surface charges and sizes, which ideallyshould be independent of the addition of QDs in the PFCphase.69 The mean size of (CF3(CF2)3)3N droplets was found todepend on the type of fluorosurfactant, while C6F14 droplet sizeswere similar for both fluorosurfactants. As expected, the dropletsmade with the anionic fluorosurfactant were highly negativelycharged in comparison to those made with the nonionic fluoro-surfactant. In all cases, the mean sizes and surface charges of the

resulting nanoscale, QD-loaded PFC droplets were similar totheir unloaded PFC droplet analogues, demonstrating that theintegration of QDs into PFC droplets did not affect these PFCdroplet properties (Figure 3).This method of silica-coated QD miscibilization into PFCs

may be easily applied to other silica-coated imaging and therapyNPs (e.g., iron oxide,56,57,60 gold nanorods (NRs),47,54 andmesoporous silica53,55,58,59 NPs). Integrating different NP-PFCcombinations together is a simple and efficient method toexpand the utility and function of current clinical imaging ortherapeutic agents.70 To show the ease of translation of NPmiscibilization into PFCs using the silica interface, the sameprocedure for QD incorporation into the C6F14 droplets wasaccomplished using silica-coated Au NRs. Hydrophilic Au NRs(with lengths of ∼35 nm and diameters of ∼10 nm) cappedwith cetyltrimethyl ammonium bromide were synthesized46

and coated with a ∼15-nm-thick mesoporous silica shell viadirect deposition,47 followed by fluorination (Supporting In-formation, Figure S3a,b). Similar to the QD integration, despitethe difference in size and shape of the Au NRs, the mesoporousmorphology of the silica layer, and the different depositionmethod for the silica coating, the Au NRs could be loaded intoC6F14 for the formation of Au NR-incorporated C6F14 droplets(Supporting Information, Figure S3c).Optical Assessment of QD-PFC Droplets in Cells in

Vitro. To assess QDs as an effective and robust tool for theassessment of PFC droplet interaction with cells, macrophage(i.e., RAW264.7) cells were selected as the model target cellfor these initial investigations.62,71 As macrophages canrapidly phagocytose anionic nanoscale agents,16,42 this cellmodel can cover the extreme ends of potential targeting

Figure 3. (CF3(CF2)3)3N andC6F14 droplets, stabilized by anionic (A)and nonionic (N) fluorosurfactants, with and without QD incorpora-tion: (a) mean droplet size and (b) surface charge.

Figure 4. Fluorescence microscopy images of fixed RAW264.7 cells(cell membrane stained green and nucleus stained blue): (a) controlcells (no QD�C6F14 droplets) and (b) cells incubated with QD�C6F14droplets (orange) for 2 h at 37 �C.

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scenarios (e.g., the best case where cells are maximally loadedwith anionic QD-PFC droplets in comparison to the casewhere control cells are unloaded giving only backgroundfluorescence signals). Also, since the reduction of PFCdroplet uptake by macrophage cells is a good indicator ofincreased in vivo circulation time, this cell model can helppredict the likelihood of successful passive targeting ofdifferent kinds of PFC droplets to tumors in vivo.16

As expected, the high resolution provided by optical micro-scopy can allow the spatial distribution of QD�PFC dropletslocalized in cells to be directly imaged for qualitative evaluation ofPFC droplet loading. For example, after a 2 h incubation of theanionic, QD-labeled, C6F14 droplets with RAW264.7 cells at37 �C, QD�C6F14 droplets internalized into the cells could beimaged using fluorescence microscopy (Figure 4b). QD�C6F14droplets (orange) were observed to be loaded into cells, within

the cell membrane (stained green) and outside the nucleus(stained blue). (The nonincubated control cells are shown forcomparison in Figure 4a.)The internalization of the QD�C6F14 droplets into the cells

was confirmed using transmission electron microscopy (TEM).As expected for phagocytic cells, the TEM images show thepresence of more vacuoles within the cytoplasm of the cellincubated with the QD�C6F14 droplets (Figure 5b) as com-pared to the control cells (Figure 5a).72 The presence of thesilica-coated QDs within these vacuoles was observed by TEM athigher magnifications (Figure 5c), indicating that these vacuolescontained the uptakenQD-incorporated C6F14 droplets. Energy-dispersive X-ray spectroscopy microanalysis of the electron-dense regions at the edges of the vacuoles showed increasedrelative concentrations of elemental Si (from the silica coating onthe QDs), Zn, Se, S, and Cd (from the QDs) in the QD-incorporated PFC loaded cells (Supporting Information, FigureS4b) as compared to the unloaded cells (Supporting Informa-tion, Figure S4a). Cu (from grid) and O and C (from the resin)were observed in both the unlabeled and labeled cells. The size ofthe vacuoles in the TEM images was observed to be larger thanthe droplets measured byDLS prior to incubation. Thismay havebeen due to temperature-induced droplet instability duringpreparation of the TEM specimen or to droplet growth withinthe vacuoles after incubation with the cells.One objective of the QD label is to help predict how different

PFC droplet formulations can interact with target and nontargetcells in vivo. Since the uptake of PFC droplets into cells is knownto be highly dependent on their surface properties,16,42 wedemonstrate that the QD labels can be used to differentiate thetime-dependent loading characteristics of anionic versus non-ionic C6F14 droplets using flow cytometry.62,71 Droplets stabi-lized by anionic fluorosurfactants should enhance phagocytosisinto macrophage cells, while the ethoxylated nonionic fluoro-surfactant headgroup should reduce their uptake.16,42 For thisdemonstration, QD�C6F14 droplets were selected becausesimilar mean droplet sizes were produced using the two differentfluorosurfactants (Figure 3a). As expected, uptake of QD�C6F14droplets stabilized by nonionic fluorosurfactants was significantlylower (<50%) than droplets stabilized by anionic fluorosurfac-tants over the 4 h incubation time (Figure 6), with uptake ofanionic QD�C6F14 droplets occurring after little as 5�15min ofincubation with the cells. Anionic C6F14 droplets (with no QDs)incubated with cells remained at baseline fluorescence over the 4h incubation time.

Figure 5. TEM images of fixed RAW264.7 cells: (a) control cells;(b) cells after a 2 h incubation with QD-C6F14 droplets at 37 �C; and(c) higher magnification of vacuoles in cells loaded with QD-C6F14droplets, showing the presence of QDs.

Figure 6. Uptake of QD�C6F14 droplets into RAW264.7 cells overtime as measured by flow cytometry, demonstrating that the moreeffective uptake of anionic compared to nonionic QD�C6F14 dropletscan be assessed using the QD label.

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The same analysis can be easily conducted on different PFC�fluorosurfactant combinations and on different target cells. Forexample, in spite of the different final droplet sizes and PFCboiling point of (CF3(CF2)3)3N droplets, their in vitro perfor-mance was similar to that of the QD�C6F14 droplets,(Supporting Information, Figure S5). This behavior was similarto other QD-incorporated PFC droplets tested. For nonphago-cytic human breast carcinoma (MDA-MB231) cells, of interest asa potential in vivo target for breast cancer,73,74 anionicQD�C6F14 droplets were uptaken in cancer cells after 1 h ofincubation via nonspecific endocytosis,75 while no measurableuptake was observed for nonionic QD�C6F14 droplets, evenafter 24 h incubation (Supporting Information, Figure S6).These types of simple and rapid optical screening methods basedon QD labeling of PFC droplets can establish baseline andselective uptake of different PFC droplet formulations with targetand nontarget cells prior to more time-consuming and expensivein vivo animal experiments.QD�PFC Droplets for Optical Assessment of PFC Droplet

Biodistribution in Histopathology. To test that the QDsintegrated in PFC droplets can be used to independently validatethe spatial distribution of PFC droplets in preclinical animalmodels, histology tissue sections (i.e., of the heart, lungs, liver,kidneys, and spleen) postsacrifice were imaged using fluores-cence microscopy after injection into rabbits. For this study,healthy New Zealand white rabbits were injected with 1.5 mL/kgof the anionic QD�C6F14 droplets via the cannulated ear vein.C6F14 droplets were selected for their potential as multifunc-tional targeted medical imaging contrast agents,9,10,12 and theanionic fluorosurfactant was selected in order to target macro-phage uptake in the Kupffer cells in the liver.76 Twenty-fourhours after injection, the rabbit was sacrificed, and the organswere removed and sectioned. Using excitation wavelengths of425�475 nm in combination with a low-pass 525 nm emission

filter and a color CCD camera, the QDs were readily identifiedusing fluorescence microscopy on unstained tissue sections ofthe liver, spleen, and lung. As an example of the type of imagesobtained, a typical section of a rabbit liver is shown (Figure 7). Ahematoxylin and eosin (H&E) stained section of the liver showsthe normal liver microstructure (Figure 7a), while an adjacentunstained section of the same area of the liver clearly shows theaccumulation of the QD�PFC agent under fluorescence micro-scopy (Figure 7b). A magnified view of an unstained section ofthe liver shows that the QDs accumulated within Kupffer cells inthe hepatic sinusoids (Figure 7c). These results demonstrate thatQD fluorescence can be maintained during fixing and standardchemical histological preparation procedures and that the QDscould be easily distinguished from the background autofluores-cence (green) of the surrounding tissue. Furthermore, the QDscan allow the fate of PFC droplets to be detected at extremelyhigh resolutions, at concentrations too low to be detected usingother clinical in vivo imagingmethods. For clinical translation, theQD tags can be removed from the PFC agent to preserve itsin vivo properties as determined by its formulation. In vitro andin vivo assessment of QD�PFC droplet localization in cells usingmedical imaging (e.g., CT, MRI, and ultrasound) is the subject offurther study. In the future, QD labels can be used to rapidlyverify the fate of the PFC droplets in vivo and help establish thein vivo targeting and clearance efficacy of different PFC dropletsfor potential clinical translation.

’CONCLUSION

New PFC droplet-based contrast agents designed for targetedclinical imaging can be assessed early in their development athigh resolution and sensitivity using in vitro optical imaging incombination with integrated QDs. This paper demonstrates asimple and robust method of miscibilizing silica-coated QDs intoPFCs, allowing optical evaluation of different PFC droplets withcells via the QD label. The method of integrating QDs into PFCdroplets can also be applied to other silica-coated, imaging andtherapy NPs. The integration of QDs did not affect the mean sizeor surface charge of the PFC droplets, and QDs could be used toevaluate the time-dependent uptake of different PFC droplets invitro using flow cytometry. For example, we confirmed thatanionic PFC droplets are taken up more rapidly than nonionicPFC droplets in macrophage cells and cancer cells. The QDscould also be used to track PFC droplet fate after in vivo injectionin a rabbit model via fluorescence microscopy of unstained tissuesections in histology. This work reveals the potential of using QDlabels for rapid, preclinical, optical assessment of different PFCdroplet formulations for their future translation into patients.

’ASSOCIATED CONTENT

bS Supporting Information. Optical characterization ofsilica-coated CdSe/ZnS QDs; microscopy of micrometer-scaleQD�C6F14 droplets; dispersion of Au NRs in C6F14 droplets;EDX microanalysis of QD�C6F14 droplets in cells; uptake ofQD�((CF3(CF2)3)3N droplets in RAW264.7 cells; and uptakeof QD�C6F14 droplets in MDA-MB231 cells. This material isavailable free of charge via the Internet at http://pubs.acs.org.

’AUTHOR INFORMATION

Corresponding Author*E-mail: matsuura@sri.utoronto.ca.

Figure 7. Microscope images of liver sections from a 3.27 kg rabbit,sacrificed 24 h after injection: (a) light microscope image of an H&Estained section; (b) fluorescent microscope image of an adjacentunstained liver section, showing the fluorescence of the QDs (orange)in the surrounding autofluorescence of the tissue (green); (c) magnifiedview of QDs in cells within the hepatic sinusoids.

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’ACKNOWLEDGMENT

Authors would like to thank Drs. James Booth and GregCzarnota for providing the RAW264.7 and MDA-MB231 cells,respectively, Siqi Zhu, Farnaz Niroui, and Grace Belayneh forassistance with the cell microscopy and flow cytometry, Dr.Joydeep Chaudhuri and Ross Williams for assistance with theanimal studies andmicroscopy of the tissue sections, and Dr. J. A.Rowlands for material support. This study was supported, in part,by the CIHR Excellence in Radiation Research for the 21stcentury (EIRR21) Research Training Program, the OntarioInstitute for Cancer Research, the Ontario Research Fund-Research Excellence Program, and the FY07 Department ofDefense Breast Cancer Research Program Concept Award(BC075873).

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