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Microvessels-on-a-Chip to Assess Targeted Ultrasound-Assisted Drug Delivery Yoonjee C Park, Chentian Zhang, Sudong Kim, Graciela Mohamedi, Carl Beigie, Jon O Nagy, R. Glynn Holt, Robin O. Cleveland, Noo Li Jeon, and Joyce Y Wong ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.6b09071 • Publication Date (Web): 26 Oct 2016 Downloaded from http://pubs.acs.org on October 26, 2016

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ACS Applied Materials & Interfaces

Microvessels-on-a-Chip to Assess Targeted Ultrasound-Assisted Drug Delivery

Yoonjee C. Park#1,7, Chentian Zhang#1, Sudong Kim2, Graciela Mohamedi1, Carl Beigie1, Jon O. Nagy3, R. Glynn Holt4, Robin O. Cleveland5, Noo Li Jeon2, Joyce Y. Wong*1,6 1

Department of Biomedical Engineering, Boston University, 44 Cummington Mall, Boston, MA 02215, USA 2 School of Mechanical and Aerospace Engineering, Seoul National University, Seoul,151-744, Korea 3 NanoValent Pharmaceuticals, Inc., 910 Technology Boulevard, Suite G, Bozeman, Montana 59718, USA 4 Department of Mechanical Engineering, Boston University, Boston, MA 02118, USA 5 Department of Engineering Science, Institute of Biomedical Engineering, University of Oxford, Old Road Campus Research Building, Oxford, OX3 7DQ, UK. 6 Division of Materials Science and Engineering, Boston University, Boston MA 02215, USA 7 Current Address: Department of Biomedical, Chemical, & Environmental Engineering, University of Cincinnati, 2901 Woodside Drive, Cincinnati, OH 45221, USA # contributed equally as first authors *Author for Correspondence: Email: [email protected]

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Abstract Microbubbles have been used in ultrasound-assisted drug delivery to help target solid tumors via blood vessels in vivo; however, studies to understand the phenomena at the cellular level and to optimize parameters for ultrasound or microbubbles in vivo are challenging and expensive to perform. Here, we utilize microfluidic microvessels-on-a-chip that enable visualization of microbubble/ultrasounddependent drug delivery to microvasculature. When exposed to pulsed ultrasound, microbubbles perfused through microvessels-on-a-chip were observed to stably oscillate. Minimal cellular damage was observed for both microbubbles and untargeted doxorubicin-encapsulating liposomes (DOX-liposomes) perfused through chip microvessels. In contrast, passive and ultrasound-assisted perfusion of integrin-targeted DOX-liposomes induced cytotoxicity, which was only significantly enhanced for ultrasound-assisted perfusion when microbubbles were co-perfused. These results suggest that stably oscillating microbubbles enhance targeted DOX-liposome internalization/cytotoxicity largely by stimulating integrin receptor endocytosis. Furthermore, our study demonstrates the utility of our microvessels-on-a-chip as a screening platform for optimizing drug dosage, targeting ligands and drugs.

Keywords: anti-angiogenesis, targeted drug delivery, focused-ultrasound, microvasculature-ona-chip, ultrasound-assisted drug delivery


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Introduction Microbubbles are increasingly being explored for ultrasound-assisted drug/RNA/plasmid DNA delivery to solid tumors through their microvasculature. Mechanistically, microbubbles have been used to directly deliver therapeutic molecules1, or to permeabilize cells in specific locations to increase uptake of therapeutic molecules delivered by other means.2-4 In the latter case, animal studies were used to show ultrasound-assisted microbubble-mediated drug or gene delivery across the blood-brain barrier into malignant tissue.2, 5 While these studies were able to experimentally establish the acoustic power and frequency required to deliver ligands across the blood-brain barrier in vivo, understanding the phenomena of which insonated microbubbles mediate this effect at the cellular level was not explained. Depending on acoustic pressure amplitude, microbubbles undergo two different cavitation modes. In the stable cavitation mode, occurring at low acoustic pressure amplitudes, microbubbles oscillate at harmonic or subharmonic frequencies and thus remain at the same equilibrium size for many acoustic cycles.6-7 In the inertial cavitation mode, occurring at high acoustic pressure amplitudes, microbubbles violently collapse, accompanied by shock wave emission. Each mode has been reported to induce specific bio-effects. Stable microbubble cavitation increases vascular permeability by producing mild microstreaming mechanical forces, which can either facilitate endocytotic uptake or mildly disrupt vascular endothelial integrity.8-11 In contrast, inertial cavitation is much more violent, and can cause transient membrane ruptures, possible cell apoptosis, and tissue injury.12-13 In this study, we have created a novel microvascular bed on a microfluidic chip (“microvesselson-a-chip”) to mimic capillary beds in vivo and used it to study the phenomena underlying ultrasoundassisted microbubble-dependent drug delivery. A passive cavitation detection (PCD) system, which recorded acoustic emissions from bubbles, was implemented to assess whether microbubbles in



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microvessels exhibit stable or inertial cavitation, and fluorescence methods were developed to assess effects of targeting and microbubble cavitation on uptake of liposomal-doxorubicin (DOX) by endothelial cells comprising microvessel walls.

Liposomal-DOX delivery was chosen for these developmental

studies for its clinical relevance14-15 and because its cytotoxic effects allowed us to gauge conditional efficacy. We employ ultrasound parameters that result in stable cavitation in chip microvessels, and show that stable cavitation is sufficient to enhance uptake of DOX-encapsulated liposomes when targeted to integrins expressed on microvessel endothelium. Our results further demonstrate that uptake and/or binding of liposomal-DOX by microvessels does not occur when they are loaded with non-targeted liposomal-DOX, even when subjected to ultrasound/microbubbles. Finally, DOX cytotoxicity studies determined that both loss of cellular integrity and cytotoxicity were greatest when microvessels loaded with microbubbles and targeted DOX-liposomes were subjected to low-energy ultrasound to stably cavitate the microbubbles. Together, these results demonstrate the potential utility of “microvessels-on-achip” in optimizing ultrasound-assisted microbubble-dependent delivery for specific drugs, a process that would be considerably more expensive and technically difficult to execute and interpret if carried out in vivo.

Results 1.

Microvessels formed in vitro allow microbubble and liposomal-DOX perfusion


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Figure 1. “Microvessels on a chip” (A) Photograph and schematic figure of microvessel-on-a-chip. The inset is a close-up of the channels into which cells were seeded and vessels were to form (B) Images of microvessel formation from Day 1 t

o Day 4 in the fibrin matrix-containing center channel. (C)



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Microbubbles in microvessels perfused with 1 um bubbles. Arrows show microbubbles that exited the microvessels. (D) Hoechst 33342 DAPI nuclear staining of the microvascular network depicted in (C).

As shown in Kim et al., vascular networks grown in the microfluidic chip provided open-ended 3D intact microvessels16 that appear similar to angiogenic microvessels17. This previous work demonstrated that endothelial cells display continuous cell–cell tight junctions as shown by the expression of the adherens junction proteins, VE-Cadherin, β-catenin, Claudin 5 and ZO-1.16 The structural stability of the microvasculature was observed by perfusing microbubbles (average diameter 1 µm) or nontargeted DOX liposomes (average diameter 78 nm) using hydrostatic pressure. In these experiments, the flow velocity through the microvessel lumen was approximately 0.03 to 0.1 mm/s depending on microvessel diameter (39.9 ± 8.58 µm, S.D), which is similar to values reported for flow velocity of red blood cells in ~40 µm diameter angiogenic vessels .18

Perfused microbubbles were observed in

microvessel lumens (Figure 1B), and in the outflow channel (arrows), indicating maintenance of vessel structure. We determined the cell-to-bubble ratio (nuclei count / bubble count, Figures 1C, 1D) to ensure consistency, which is approximately 1:13 when ~108/mL microbubbles were introduced in microvessels. Results also show the microvessels are impermeable to fluorescently labeled non-targeted DOX liposomes (Figure 2). Together these data indicate that the walls of microvessels-on-a-chip are openended, consist of tightly connected endothelial cells similar to capillaries, are impenetrable to nano-sized particles (~78 nm), and can accommodate physiological flow rates.

2. Targeted DOX-liposomes specifically bind to microvessels under flow We next examined the effect of endothelial cell-targeting on retention of liposomes by the walls of chip microvessels (Figure 2). The brightness of the fluorescence signal was expressed using a color map programmed in ImageJ, called Red Hot, which makes: low brightness intensity red, the intermediate brightness is yellow, and the brightest signal is white (as shown in Figure 2). We have analyzed 5 samples


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for each condition using ImageJ and obtained the ‘mean grayscale brightness intensity’. The. Quantitative fluorescent intensity values are 244. 5 (st devstd =4.95) for the non-targeted DOX liposomes and 289.1 (st devstd=7.89) for the targeted DOX liposomes. The results clearly show that non-specific binding of non-targeted DOX-liposomes to microvessel walls is negligible compared to that seen with targeted DOX-liposomes, an indication of effective RGD-dependent integrin targeting.

Figure 2. RGD-dependent targeting of DOX-loaded liposomes to chip microvessels. Non-targeted and integrin-targeted liposomes were perfused into chip microvessels. Images were taken under TRITC filter before and after washing to determine DOX fluorescence. Fluorescence brightness was expressed using a color map (right). Images are representative of 3 experiments.

3.

Microbubble cavitation in microvessel lumens

Samples were exposed to 1.1 MHz ultrasound with a peak negative pressure 800 kPa at a pulse repetition frequency of 100 Hz and a duty cycle of 0.1% for a duration of 2 min (see Figure 7). In order



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to understand how ultrasound-microbubble (MB) interaction might affect delivery of DOX by nontargeted/targeted liposomes, the oscillatory behavior of MBs in microvessels-on-a-chip was assessed by analyzing acoustic emission waveforms acquired from the passive cavitation detector (PCD) hydrophone over 2 min of insonation (see Figure 7). Figure 3 shows the high-pass (fcutoff = 1.5 MHz) filtered frequency spectra obtained at commencement of insonation and just prior to its termination at 2 min. Microvessel preparations receiving MB alone (n = 2), targeted DOX-liposomes alone (n = 3), and targeted DOX-liposomes+MB (n = 3) were examined. Prominent 2nd harmonic (2.2 MHz) peaks were observed for MB alone and MB+targeted liposomes. These peaks were not observed in the absence of MB (targeted DOX-liposome alone group), indicating they are characteristic of intact MB. To determine if inertial cavitation was present, we compared the integrated broadband signal between harmonic peaks (see Methods)

19

with that obtained during the last 10 cycles just prior to insonation cessation at 2 min.

No significant differences were observed in all three conditions (see Supporting Supporing Info) . These emission results suggest that bubble activity in this study is mainly stable cavitation, not inertial cavitation, as would be expected for the 800 kPa peak focal pressure used.19 There was, however, a significant decline in peak harmonic amplitudes in the two preparations containing microbubbles over the two minute experiment duration, implying a significant number of bubbles were destroyed even in the absence of significant inertial cavitation.


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Figure 3. Acoustic emission Fast Fourier Transform (FFT) data from (A) Targeted liposomes alone (B) Microbubbles alone and (C) Targeted liposomes + microbubbles. Left column plots are FFTs of the first 10 cycle pulse of insonation and right column plots are FFTs of the final 10 cycle pulse of insonation (1.1 MHz, 2 min., 800 kPa peak negative pressure).

4. Microbubble cavitation permeabilizes microvessels to targeted liposomal-DOX In order to observe the interactive effects of cavitation and DOX-mediated toxicity on microvessel integrity, we compared microvessel morphology 24 hr after they were insonated with targeted DOX-liposomes, either in the presence or absence of MB (Figure 4). Although noticeable fluorescence indicated that significant DOX-liposomes became microvessel-associated with ultrasound alone, the microvessels remained intact (bright-field), with little or no accumulation of DOX in the fibrin



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gel outside the vessels. In contrast, insonation when MB were added to the liposomes (microbubble cavitation) resulted in disruption of microvessel walls (black arrows, bright-field image) and significant DOX accumulation in the surrounding fibrin gel (white arrows, TRITC-filtered image). Numerous bright yellow spots were also observed along the microvessel walls under the TRITC filter when both targeted DOX-liposomes and MB were present, indicating a greater accumulation delivery of cytotoxic DOX to microvessel walls than in the absence of cavitating MB. A similar study was also performed with NTD (non-targeted liposomal-DOX) to reveal whether the DOX accumulation in the fibrin gel is due to vascular permeation or exocytosis of DOX after endocytosis. Insonation in either the presence or absence of MB did not diminish vessel integrity (Figure 5C) and, consistent with this finding, did not generate a significant fluorescent signal in the surrounding fibrin gel (data not shown). Thus, the results indicate that DOX penetration through gross vascular permeation is not the dominant effect of insonation in this study. Instead, receptor-mediated DOX endocytosis appears to have been enhanced by low power microbubble cavitation. Further experiments relating to DOX cytotoxic efficacy on the vessels described below also support this conclusion.


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Figure 4. Ultrasound and microbubbles increase toxicity of DOX-loaded liposomes targeted to integrins. Microvessels were perfused with integrin-targeted liposomes alone (A) or integrin-targeted liposomes + microbubbles (B), then washed and insonated (1.1 MHz, 2 min., 800 kPa peak negative pressure). Bright field (top) and TRITC fluorescent (bottom) images were taken after 24 hr. Black arrows in the bright field image in panel B show cells extruded from nearby vessels. White arrows in TRITC image in panel B show liposomes that have penetrated the external vessel walls.

5. Cavitation increases cytotoxicity of liposomal-DOX on microvessels Because the results in Figure 4 showed a MB-dependence of ultrasound-facilitated endothelial disruption, we next wanted to correlate this effect with cytotoxicity analysis. Ten different combinations



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of variables were tested for cytotoxicity, comparing the effects of: insonation on or off, the presence or absence of MB, the presence or absence of DOX liposomes, and targeted vs. untargeted DOX-liposomes. We first evaluated cytotoxic effects of MB with and without ultrasound. Figure 5(A) indicates that the insonation regime employed here has no statistically significant effect on cell mortality in the presence of MB alone. Figure 5B shows that the cells appear healthy and similar to what is seen in control samples (data not shown).

Thus in these experiments, the absence and presence of insonation

resulted in the death of 1.8±0.2% (mean± S.D.) and 3.4±1.8% of the microvessel endothelial cells, respectively. The next group of experiments examined cytotoxicity when non-targeted DOX-liposomes were subjected to insonation in the absence or presence of MB. Visual examination of the photomicrograph (Figure 5C) shows that non-targeted liposomes are not cytotoxic in the presence of potentially damaging insonated microbubbles. The quantified results in Figure 5A confirm this visual impression. Mortality rates induced by non-targeted DOX-liposomes without and with ultrasound were 4.2 and 5.0±2.2%, respectively; those induced by non-targeted DOX liposomes + MB without and with ultrasound were 4.3±0.8% and 5.7±3.9%, respectively. Thus, although the data indicate cytotoxicity trends higher when non-targeted DOX-liposomes and cavitating microbubbles are present (compared to the baseline condition with neither liposomes nor MB), the effect was not statistically significant. In contrast, there was a highly significant increase in cytotoxicity when the same 4-way experimental design was repeated with targeted DOX-liposomes. It can be seen in Fig 5A that there was roughly 30% cell mortality when targeted DOX-liposomes were tested without MB, regardless of whether the microvessel preparation was subjected to insonation (28.6±5.7% without ultrasound, 27.2±4.9% with ultrasound), and, as the images in Fig 5D show, a similar mortality rate was observed when microvessel preparations were incubated with targeted DOX-liposomes and MB but not insonated (32.0±7.2%). Consistent with the data in Figure 5A, by far the greatest cytotoxicity was observed when microvessel preparations perfused with targeted DOX-liposomes and MB were subjected to ultrasound (Figure 5E).


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Mortality in these experiments averaged 52±6.7%, and was significantly greater (p 99%) and 15 mol% of 1,2-distearoyl-sn-phosphoethanolamine-PEG5000 (m-PEG5000-DSPE, purity >99%) (Avanti Polar Lipids) were used as microbubble shell materials. The mixture in chloroform (SigmaAldrich, purity >99%) was evaporated in a chemical hood, and the dry film was hydrated with a 10/10/80 (v/v/v) solution, which consisted of 10% glycerin, 10% propylene glycol (Sigma-Aldrich, St. Louis, MO, purity > 99%), and 80% water, resulting in a total concentration of the lipid of 5.32 µmol/mL. The mixture was tip-sonicated for 10 min until the dispersion became clear (Branson 450 Sonifier 400W). 200 µL of the lipid dispersion was added in a septum vial, and the head space air was replaced with decafluorobutane gas (Synquest Laboratories, Alachua, FL) to make microbubbles. The gas-filled vial was shaken by Vialmix (Lantheus Medical Imaging, North Billerica, MA) to generate microbubbles, and the average size of the microbubbles was measured via standard bright field microscopy images. The microbubble dispersion was diluted 1:10 in an endothelial growth medium (EGM-2) (Lonza, Walkersville MD), and then loaded in two medium reservoirs on the side where microvessels initiate (Figure 1A).

3. Generation of non-targeted and targeted doxorubicin (DOX)–containing liposomes Both targeted and non-targeted polymerized liposomes were used to encapsulate DOX. The preparation protocols for polymerized liposomes were adapted from Federman et al.26 Non-targeted DOX-liposomes were prepared from (5’-hydroxy-3’-oxypentyl)-10-12-pentacosadiynamide (h-PEG1PCDA) (NanoValent Pharmaceuticals, Inc. Bozeman, MT), L-α-phosphatidylcholine, hydrogenated Soy (hydro soy PC), 1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(polyethylene glycol)2000] (m-PEG2000-DSPE) and cholesterol (Avanti Polar Lipids, Alabaster, AL, purity >99%), at a molar proportion of 15:47:32:6, according to the method previously described.22 The synthetic lipid, h-PEG1PCDA, was purified by silica gel chromatography and recrystallization to give a material that is analytically pure >99% by elemental analysis. For targeted DOX-liposome formulation, 1,2-distearoylsn-glycero-3-phosphoethanolamine-N-[biotinyl(polyethylene

glycol)-2000]


(biotin-PEG2000-DSPE)

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(Avanti Polar Lipids, Alabaster, AL, purity >99%) was used to make biotin functionalized vesicles. Hydro soy PC, cholesterol, biotin-PEG2000-DSPE, and m-PEG2000-DSPE, at a molar proportion of 15:47:32:4.5:1.5, were mixed and evaporated in vacuo (0.002 torr) to a film. 155 mM ammonium sulfate (pH 5.5) was added to the films so as to give a 10 mg/mL total lipid and cholesterol suspension. The suspension was heated via sonication between 70 and 80⁰C with a tip sonicator (Fisher Sonic Dismembrator model 300) for 5 min. The resulting slightly milky solution was then extruded through a stacked polycarbonate membrane (100 nm), eleven times, with a dual syringe extruder (LiposoFast-Basic, Avestin, Inc., Ottawa, ON, Canada), heated at 65⁰C. The nearly clear liposome solutions were cooled to 5⁰C for 12 hr. After warming to ambient temperature, liposomes were polymerized by UV light irradiation (254 nm) with a SpectrolinkerXL-1000 UV Crosslinker (Spectronics Corporation, Westbury, NY) for 10 min. The resulting blue polymerized liposomes were heated to 65⁰C for 5 min to convert them to the red form. The colored solutions were syringe-filtered through 0.2 µm cellulose acetate filters in order to remove trace insoluble contaminants. The ammonium sulfate-containing polymerized liposomes were passed over a G50 Sephadex column (washed with PBS buffer, pH 7.4) to exchange the external buffer. The liposomes were then incubated (65⁰C for 20 min) with doxorubicin HCl (LC Laboratories, Woburn, MA) at a concentration of 0.5 mg of DOX to 1 mg of liposomes. The free doxorubicin was removed by mixing the products for 5 min with an anionic exchange resin (Bio-Rex 70, Bio-Rad Inc) in a ratio of 7 µg of DOX to 1 µL of packed resin. DOX-encapsulating liposomes were separated from the resin by filtering through Pierce Spin Cups. The average particle size measurements were obtained on a Zetasizer Nano S (Malvern Inst.), in a solution of 10 mM sodium chloride. Targeted DOX-encapsulating liposomes were synthesized as follows. Biotinylated Arg–Gly–Tyr (RGD) (Mimotopes, Australia) was used as targeting ligand. 400 µL of washed DOX-encapsulating liposomes (~1.9 x 1013 particles) was incubated with 8 µL of 0.5 mg/mL neutravidin (Life Technologies, Grand Island, NY), for 5 min and ultra-centrifuged in a centrifugal filter (Centricon Plus 70, 100 kDa,



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Millipore, Billerica, MA) at 3500 g for 10 min to remove free neutravidin. Neutravidin-containing liposomes remaining in the filter were recovered by centrifuging at 1000 g for 2 min. 8 µL of 1 mg/mL Biotinylated RGD peptide was then added to the liposome suspension, followed by a 30 min incubation period. Thereafter, the liposomes were dialyzed for 24 hr to remove unincorporated peptide (Schematics in Figure 6). When introduced in microvessels, 0.16 mM DOX in EGM-2 media was used.

Figure 6. Schematic of targeted DOX-loaded liposomes and microbubbles used in this study.

4. Generating Microvessels on-a-Chip Capillary-like microvessels were generated in a microfluidic device according to the method of Kim et al.16, with optimization for ultrasound setup (Figure 1A). Lung fibroblasts (LF) were seeded in the rightmost channel in Figure 1B, while the center channel, into which the vessels were to grow, was filled with a fibrin gel. Human umbilical vein endothelial cells (HUVEC) were introduced into the medium channel contralateral to the fibroblast channel, and grew into the center channel in response to factors secreted by the fibroblasts. Briefly, microfluidic devices were fabricated using soft lithography. Polydimethylsiloxane (PDMS, Sylgard 184, Dow Corning) was cured against a microfabricated mold overnight at 80⁰C. The molded PDMS piece was punched out for hydrogel injection ports and reservoirs for cell culture media (Figure 1A). 3.5g of PDMS was cured in a 100 mm Petri dish to create a thin PDMS sheet (~150 µm).


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The molded piece and the thin sheet were treated with air plasma for 45 sec and bonded together irreversibly. The PDMS devices were kept at 80⁰C for at least 24 hr and sterilized by UV irradiation for 30 min before each experiment.16 Lung fibroblasts (LF) (Primary cells, Lonza) were cultured in FGM media (Lonza), harvested and resuspended in FGM. A fibrinogen solution containing 2.5 mg/ml bovine fibrinogen (Sigma), and 0.15 unit/ml aprotinin (Sigma) was combined with the lung fibroblast cell suspension at a concentration of 13x106 cells/ml. The cell/fibrinogen mixture was then mixed with thrombin (1 unit/ml, Sigma), and immediately injected into a single cell culture channel (rightmost channel in Figure 1A). The central channel and leftmost channel were filled with 2.5 mg/ml acellular fibrin solution (0.15 unit/ml aprotinin, 1 unit/ml thrombin) and left to clot at 37⁰C for 7 min. After gelation, EGM-2 medium (Lonza) was loaded into one side of the reservoirs connected to the cell culture medium channel and vacuum was applied on the other side to fill the medium channels. Then the four reservoirs were uniformly loaded with EGM-2 medium. Devices were incubated for 24 hr at 37⁰C and 5% CO2 to allow air bubbles captured between the gel–medium interface to dissolve and fibroblasts to adhere and spread out within the fibrin matrix. Thereafter, human umbilical vein endothelial cells (HUVEC, Lonza), previously cultured in EGM-2 medium, were harvested and resuspended in EGM-2 medium at a concentration of 5x106 cell/ml. Medium reservoirs were emptied, and 4 µl of HUVEC suspension was introduced into the medium channel contralateral to the lung fibroblast seeding (Figure 1A). Devices were then tilted by 90° and incubated at 37⁰C and 5% CO2 for 40 min to allow cell attachment. After cell attachment, fresh EGM-2 medium was gently added to reservoirs, and devices were incubated at 37⁰C for several days to allow endothelial cells to grow into microvessels. The medium was changed after 3 days. Fully perfusable vessels were usually ready to use after 5 ~ 6 days (Figure 1B).

5.

Introducing microbubbles and DOX-liposomes in microvessels-on-a-chip



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Microbubbles or DOX liposomes were introduced in the microvessels using hydrostatic pressure. The flow velocity through the microvessel lumen caused from hydrostatic pressure was approximately measured by analyzing captured videos. Flow stopped when liquid levels on inlet and outlet sides were even, which usually occurring within 5 min. Either non-targeted or targeted fluorescently labeled DOXliposomes were perfused through microvessels in the device, followed by wash with media to remove unbound liposomes.

6. Ultrasound setup and insonation protocol The experimental arrangement for ultrasound exposure is shown in Figure 7. A single-element spherically focused piezoceramic transducer (H-102, Sonic Concepts, Bothell, WA), nominal center frequency of 1.1 MHz, 63-mm radius of curvature, 64-mm aperture diameter and a 20-mm hole for a monitoring transducer, was employed as the focused ultrasound source. A function generator (33120A, Agilent Technologies, Palo Alto, CA) supplied the 1.1 MHz sinusoidal signal that was amplified (55 dB, A150, ENI, Rochester, NY) before passing through a custom impedance-matching network (Sonic Concepts). The free-field acoustic pressure was determined using a pressure calibration performed in water using a calibrated PVdF needle hydrophone (75 Micron Probe, SN 1355, Precision Acoustics, Dorchester, U.K.). After a sample was introduced into the microvascular bed in the microfluidic device, the device was positioned at the focus All measurements were performed in a tank (45 cm × 45 cm × 58 cm) filled with deionized and degassed water. Insonation was carried out for 2 min with 10-cycle long pulses with a peak negative pressure of 800 kPa at a pulse repetition frequency of 100 Hz (a duty cycle of 0.1%).


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Figure 7. A schematic diagram of the experimental setup for ultrasound. The zoomed-in schematic shows a cross-section of the device. Ultrasound was applied from a focused ultrasound transducer at 1.1 MHz for 2 min with a peak negative pressure of 800 kPa and the bubble activity was detected by passive cavitation detector (hydrophone).

7. Passive cavitation detection and acoustic emission analysis Acoustic emissions consisting of the scattered primary 1.1 MHz and its harmonics plus bubble emissions were received by a 5 MHz unfocused hydrophone transducer (0.5” diameter, V309-SU Panametrics-NDT Corporation, Waltham, MA) positioned orthogonal to the insonifying transducer (Figure 7). The collected signal was then amplified (Panametrics Ultrasonic Preamp) by 34 dB and digitized and saved on a digital oscilloscope (WaveRunner 6050A, LeCroy, NY). All post-processing was conducted using purpose written MATLAB-code (R14, The MathWorks, Natick, MA). The digitized waveform was filtered using a highpass filter (created with MATLAB signal processing toolbox) with a stopband frequency of 1.5 MHz and passband of 2 MHz. The power spectrum was analyzed using the periodogram function, and the Fourier Transform was produced via the FFT function. Following the



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observation that the 4th harmonic (4.4 MHz) was consistently absent from the PCD FFT, the amplitude spectrum was integrated between 3.5MHz to 5 MHz to quantify broadband acoustic emissions specifically from inertial cavitation activity. After filtering the fundamental (1.1 MHz), 3rd and sometimes 5th harmonic peaks were observed even in the absence of MB, which was characteristic of the H-102 transducer.

8. Imaging To study the uptake of DOX-encapsulated liposomes, devices were loaded with a sample for 15 min and then washed with EGM-2 medium. Doxorubicin has intrinsic fluorescent properties.35 After a further incubation for 24 hr, bright-field and fluorescent images were acquired using an Axiovert S-100 microscope system (Carl Zeiss, Inc., Thornwood, NY) at 10x magnification equipped with a LED fluorescent light source (Spectra light engine, Lumencor, Beaverton, OR). The microscope was controlled by Metamorph imaging software (Molecular Devices, Sunnyvale, CA). The brightness of the fluorescence signal was expressed using a color map programmed in ImageJ, called Red Hot, which makes low brightness intensity red, the intermediate yellow, and the brightest white (as shown in Figure 2). For fluorescence intensity analysis, 5 samples for each condition was analyzed via ImageJ to obtain mean grayscale brightness intensity.

9. Cell mortality analysis To study cell mortality, the devices exposed to ultrasound with samples were incubated for 15 min, followed by washing with EGM-2 medium, all at 37⁰C. Thereafter they were incubated for an additional 24 hr at 37⁰C and 5% CO2 to observe cytotoxicity. Devices were then thoroughly washed twice with PBS, and cell staining dyes were loaded and incubated at room temperature for 15 min prior to imaging. The PBS (Fisher) used contains potassium phosphate monobasic (KH2PO4) at 1.06 mM, sodium


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chloride (NaCl) at 155.17 mM, and sodium phosphate dibasic (Na2HPO4-7H2O) at 2.97 mM. Cell mortality on microvessels was quantified by using two types of cell staining dyes. Hoechst 33342 (Life technologies) was used to count the total number of cells comprising the microvessels because it emits blue fluorescence when bound to dsDNA present in both live and dead cells.36 To count the number of dead cells, SYTOX® Green Dead Cell Stain (Life technologies) with excitation at 488 nm with an emission profile similar to FITC (521 nm) was used to avoid overlap with the fluorescence signal from either polymerized liposomal-DOX (excitation: 547 nm; emission: 557 nm) or DOX (excitation: 425 nm; emission: 600 nm). 30 µL of a mixture of 5 µM of Hoechst 33342 and 1 µM of SYTOX Green was added in two medium reservoirs in a device where a sample was loaded after suctioning the medium retained in the device channels. Devices were incubated at room temperature for 20 min in the dark and imaged. The number of total or dead cells were counted using ImageJ (National Institutes of Health) software. Finally cell mortality was calculated as follows: # ( )

Cell mortality for non-targeted DOX-liposomes = # ( ) x 100 (%) Cell mortality for targeted DOX-liposomes =

# ( ) # ( )

x 100

(%) For the targeted DOX-liposome group, cell mortality was calculated differently from other conditions because nuclei fragmented when cells were dead and severely damaged, resulting in an uncountable number of SYTOX green spots. Live cells were counted from Hoechst 33342 images. Because nuclei that are rounded in Hoechst 33342 images also show up in SYTOX Green images (dead), nuclei with normal shape from Hoechst 33342 images were considered as live cells. This method, however, slightly underestimates cell mortality because of cell aggregation, which makes it difficult to quantify cell number and results in an apparent decrease in the total number of cells (Figure 5A).

10. Statistical analysis



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Cell mortality data were obtained from three or more independent observations conducted on different microfluidic devices, where the value for each observation of the same microvessel perfusion condition was pooled for calculating average and standard deviation (S.D.). Quantitative data in histograms are expressed as means ± S.D. Statistical significance between different microvessel perfusion conditions was determined using one-way ANOVA followed by the Tukey-Kramer post-hoc test for multiple comparisons. A p-level of < 0.05 was considered to be significant. All analysis was done using the Matlab statistics toolbox.

Supporting Information Integrated broadband signal of acoustic emission Fast Fourier Transform (FFT) data during the last 10 cycles.

Acknowledgements We thank Stanley Heydrick for careful reading and editing of the manuscript. J.Y.W. acknowledges a Boston University College of Engineering Distinguished Faculty Fellowship.

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TOC 256x161mm (150 x 150 DPI)


Microvessels-on-a-Chip to Assess Targeted ... - ACS Publications

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