Rapid Assessment of Nanoparticle Extravasation in a Microfluidic

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Rapid Assessment of Nanoparticle Extravasation in a Microfluidic Tumor Model Mai Ngoc Vu, Pradeep Rajasekhar, Daniel Poole, Song Yang Khor, Nghia P. Truong, Cameron J. Nowell, John F. Quinn, Michael R. Whittaker, Nicholas A Veldhuis, and Thomas P. Davis ACS Appl. Nano Mater., Just Accepted Manuscript • Publication Date (Web): 29 Mar 2019 Downloaded from http://pubs.acs.org on March 29, 2019

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ACS Applied Nano Materials

Rapid Assessment of Nanoparticle Extravasation in a Microfluidic Tumor Model Mai N. Vu1,2,3,4, Pradeep Rajasekhar1,3, Daniel P. Poole1,3, Song Yang Khor1,2, Nghia P. Truong1,2, Cameron J. Nowell3, John F. Quinn1,2, Michael Whittaker1,2, Nicholas A. Veldhuis1,3,*, Thomas P. Davis1,2,5* 1Australian

Research Council Centre of Excellence in Convergent Bio-Nano Science and

Technology, Monash University, Parkville, VIC 3052, Australia; 2Drug

Delivery, Disposition and Dynamics, Monash Institute of Pharmaceutical Sciences,

Monash University, Parkville, VIC 3052, Australia; 3Drug

Discovery Biology, Monash Institute of Pharmaceutical Sciences, Monash University,

Parkville, VIC 3052, Australia; 4Department

of Pharmaceutics, Hanoi University of Pharmacy, Hanoi, 10000, Vietnam;

5Department

of Chemistry, University of Warwick, Gibbet Hill, Coventry CV4 7AL, United

Kingdom.

KEYWORDS: microfluidics, tumor, EPR, edema, TRPV4, endothelium

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ABSTRACT The development of nanoparticle-based targeted therapeutics for the treatment of cancer requires a well-defined understanding of the tumor microenvironment, which is challenging due to tumor complexity and heterogeneity. Recent advancements in three-dimensional (3D) cell models such as tumor-on-a-chip devices can overcome some of these challenges by providing co-culture in vitro systems (tumor surrounded by tubular endothelial cells) that mimic native cellular environments to accurately study the enhanced permeability and retention (EPR) potential of drug delivery systems under flow conditions. However, inducing “leaky” vasculature in endothelial cells surrounding solid tumors in microfluidic devices is not readily controllable and highly dependent on tumor cell identity. Utilizing a microfluidic tumor model (MTM) consisting of a tumor region surrounded by a 3D microvascular network, we have simulated the EPR effect by activating a known regulator of endothelial junction formation and edema: the Transient Receptor Potential Vanilloid 4 (TRPV4) ion channel, to rapidly assess extravasation and tumor accumulation of nanoparticles of different sizes and surface chemistries. Treatment with a selective TRPV4 agonist stimulated reorganization of the actin cytoskeleton and disruption of adherens junctions to provide a concentrationdependent or “tunable” leakiness, confirmed by increased tumor uptake of fluorescent dextran macromolecular tracers from the vascular channels. While this controlled 3D in vitro vascularedema system may not exemplify all of the complexities of edema mechanisms in vivo, it provides a rapid, materials-focused screening method to assess the extravasation and tumor uptake potential of nanoparticles with distinct properties. We show that the passage of nanoparticles through leaky vasculature is not solely governed by particle size but also by surface chemistry, where surface tertiary amines limit tumor cell association due to unwanted endothelial interactions.


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INTRODUCTION As a therapeutic strategy for treating tumors, drug delivery systems such as nanoparticles (NPs) routinely exploit the enhanced permeability and retention (EPR) effect for passive targeting and uptake. This requires leaky tumor vasculature to facilitate NP entry, and poor lymphatic drainage to limit NP clearance from within a tumor region1-3. Despite considerable effort in the field to utilize the EPR effect for therapeutic success, a strikingly small proportion of nanoparticle types have been reported to accumulate in solid tumor targets4-5. This may be associated with complicating factors related to the properties of drug delivery systems or tissue heterogeneity, which is influenced by cancer identity, progression and leakiness of tumor vasculature6. However, in the absence of surface targeting moieties such as small ligands or antibodies, the modification of NP physicochemical properties (e.g., surface chemistry, size, charge, or shape) continues to offer promise for influencing tumor delivery and uptake, and requires suitable systems to assess their uptake and efficacy7-9. Tumor-endothelial interactions are complex and involve many factors, including tumor type, location and stage of tumor development. Hence, no single in vitro system can recapitulate all of these factors to reproducibly and rapidly assess nanomaterial properties that may be optimal for anti-tumor therapies. Recent evidence suggests that greater physiological representation of vascular architecture and tumor microenvironments can improve the uptake, toxicity profiles or drug resistance of anticancer agents, and hence, the development of in vitro microfluidic or 3D culture tumor models for pre-clinical screening offers valuable systems for rapid assessment and optimization of drug carriers, to bridge the gap between standard in vitro approaches and in vivo studies10, 11-12. Tumor spheroids can incorporate multiple cell types and enable drug testing on primary cultures from patient-sourced tissues, but cannot be readily exposed to dynamic flow conditions to simulate the transport of NPs from the surrounding tumor vasculature13. To assess the potential for NPs to undergo extravasation, microfluidic



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devices can measure the impact of shear forces on tumor or endothelial cells and enable assessment of diffusion of particles across endothelial barriers. Systems that employ parallel straight channels inaccurately mimic the branched vasculature and variable shear forces observed in vivo, therefore leading to limitations in our understanding of the behavior and cellular interactions of NPs in complex vascular networks14-16. To overcome this, microfluidic tumor models (MTMs) have been developed as co-culture systems incorporating an interstitial space for tumor cells, surrounded by a 3D vascular network of interconnected, nonlinear microfluidic channels mapped from structures of capillaries imaged in vivo17. This provides dynamic flow conditions for nanoparticle or drug delivery, with wall shear rates that are heterogeneous across different regions of the chip, to mimic native environments and provide rapid assessment of nanoparticle uptake from microfluidic channels into the tumor space18-19. A 2 μm porous spacer at the interface between vascular and tumor compartments allows macromolecules and secreted cellular factors to freely diffuse between vasculature and tumor regions20. This provides greater physiological relevance compared standard 2D in vitro culture systems, and can be utilized for rapid preclinical screening of anti-cancer agents, to assess physicochemical properties of NPs that may influence their extravasation, EPR potential, toxicity or drug release profiles. MTMs have the potential to offer viable alternatives to animal tumor models, by avoiding ethical and practical issues, including lack of reproducibility, cost and animal cancer burden21. In this study, a Synvivo™ MTM was utilized to assess how nanoparticles of different sizes and surface chemistries pass across endothelial barriers (extravasation) and accumulate in tumor cells. These studies were performed with consistent flow (cell exposed to heterogeneous shear forces across the device) using control media conditions, or with “induced-edema” conditions, where vascular channels were perfused with GSK1016790A (GSK101), a highly selective agonist for the Transient Receptor Potential Vanilloid 4 (TRPV4)


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cation channel. TRPV4 activity is associated with barrier formation and is known to promote edema through Ca2+-mediated regulation of actin cytoskeletal and VE-cadherin junction proteins. Consequently, TRPV4-mediated loss of cell barrier integrity has been observed in multiple tissues, including vascular endothelia, lung, colon and retinal epithelia22-25 and was therefore proposed to be a viable target for controlling the leakiness of the tumor microvasculature to modulate nanoparticle EPR. Previous reports using microfluidic devices have induced leakiness in tumor endothelial cells using conditioned media from tumors. However, the precise cellular factors produced have not been accurately determined for all tumor types and more importantly, supernatants from different tumor cell sources result in different levels of permeability21. In this proof of concept study, we applied an alternative reductionist approach for rapidly assessing nanoparticle properties in a co-culture system under flow conditions. Using a microfluidic 3D culture system, we confirm that TRPV4 activity modifies endothelial cytoskeleton and junction proteins (i.e., actin and VE-cadherin), as shown by the increased presence of intercellular gaps and changes in the intracellular distribution of actin in a GSK101 concentration-dependent manner. By measuring the extravasation and accumulation of fluorescent macromolecular dyes and nanoparticles across the vascular barrier into the tumor area, we demonstrate that pharmacological control of TRPV4 in an MTM system offers an efficient, tunable method to control vascular permeability, to mimic heterogeneous leakiness of tumor vasculature, and to provide a reproducible system for identifying nanoparticle properties that are important for extravasation and tumor accumulation.



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RESULTS AND DISCUSSION Establishment of a vascularized microfluidic tumor model. A vascularized MTM has been developed and characterized, consisting of a tumor compartment and a surrounding network of vessels, separated by a 2 μm-wide porous interface (Figure 1A, B)17, 21, 26. Breast cancer MDAMB-231 cells stably expressing green fluorescent protein (GFP) were seeded in the tumor region 24 h before seeding and growing Human Umbilical Vein Endothelial Cells (HUVECs) to confluence in the vascular channels, followed by continual perfusion of endothelial cell growth media (EGM) for a further 22 h at 0.1 µL min-1. HUVECs were chosen as the primary endothelial cell line of choice for these proof of concept studies, due to their reproducibility in seeding microfluidic devices, prior tumour-on-a-chip studies27 and evidence of HUVEC TRPV4 expression, function and cell junction regulation in previous edema-focused studies24, 28.

The segmentation of vascular endothelial and tumor cells was confirmed by imaging

analysis of GFP fluorescence to confirm the absence of tumor cells within the 3D vascular network (Figure 1B, C). GFP fluorescence was also used to distinguish MDA-MB-231 from HUVECs in subsequent quantitative flow cytometry experiments. The integrity of the endothelial lining plays an important role in controlling transport of solutes and circulating cells from blood vessels into interstitial tissues29. Expression of the endothelial cell (EC) specific marker platelet-endothelial cell adhesion molecule-1 PECAM-1 (CD31) was confirmed in HUVECs cultured in the MTM system (Figure 1D). To characterize the formation of complete and intact HUVEC endothelial barrier, the cells were immunolabeled with vascular endothelial cadherin (VE-cadherin) for detection of adherens junctions (AJ) and fluorescentlylabeled phalloidin for detection of filamentous actin (F-actin). Representative images indicate that HUVECs formed a confluent, microvascular network with a tubular 3D cellular architecture and established VE-cadherin-positive adherens junctions (Figure 1E).


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TRPV4 agonist disrupts endothelial junctions and induces actin cytoskeletal reorganization. We sought to develop an in vitro vascular model that could be readily modulated by a pharmacological agent to disrupt the intercellular junctions of endothelial cells and mimic the leakiness of tumor vasculature observed in vivo. TRPV4 was hypothesized to be a suitable target due to its established role in barrier formation and edema, the availability of potent selective agonists and its known interactions with endothelial junction proteins (cadherins and -catenin)

30,

and cytoskeletal actin or tubulin proteins22-25, 31. We perfused

increasing concentrations of the TRPV4-selective agonist GSK1016790A (GSK101; 1, 3, and 10 nM) into the MTM chip for 30 min at a flow rate of 1.0 μL min-1. These concentrations were confirmed to have no significant effect on cell viability of HUVECs or MDA-MB-231 cells in the MTM system (Figure S1). To observe changes in endothelial cell junctions, cells were stained for VE-cadherin, fluorescently-labeled phalloidin and Hoechst 33342 (nucleus) and then imaged by confocal fluorescence microscopy (Figure 2A). In endothelial cells, immunofluorescent labelling of VE-Cadherin to track adherens junction formation and the use of fluorescently-labeled phalloidin to assess F-actin distribution, cell morphological changes and gap formation has been described previously, and demonstrated important roles for TRPV4 and TRPV4 endogenous mediators in cytoskeletal rearrangement3235.

In non-stimulated control conditions, F-actin microfilaments were evenly distributed across

the cells (Figure 1E) and HUVECs formed intact junctions with expression of three specific types of AJs, including linear and discontinuous junctions along cell-cell borders, and reticular AJs distributed in overlapping areas between adjacent, neighboring cells (Supplementary Figure S2)36-37.



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Figure 1. The 3D microfluidic tumor model. (A) Schematic of the MTM with vascular network (blue), porous interface (gray) and tumor area (red). Inlet and outlet ports allow cells to be maintained by consistent media perfusion. (B) Bright field image of the tumor chip with endothelial cells (HUVECs) in the vasculature, cancer cells (MDA-MB-231) in the tumor compartment, and porous architecture of 2 μm interface separating vessels and tumor area. (C) GFP-positive MDA-MB-231 cells (green channel) in the tumor area are separated from HUVECs seeded into the vascular network; all cells indicated by Hoechst 33342 nuclear stain (blue channel); scale bar = 200 μm. (D) HUVECs in the vascular network confirmed by immunolabeling with CD31 immunostaining (red channel) and Hoechst 33342 nuclear staining (blue channel); scale bar = 100 μm. (E) Vascular architecture (top and side view) in confluent HUVECs (24 h post-seeding) confirmed by labelling of the cytoskeleton (actin, green channel), junction protein (VE-cadherin, red channel), and nuclei (Hoechst 33342, blue channel). Cells maintain a tubular formation and three-dimensional lumen; scale bar = 20 μm.


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Studies in open flow systems indicate that increased shear forces lead to moderate increases in TRPV4 cell surface expression and Ca2+ flux, suggesting that under flow conditions actin organization may be maintained by TRPV4-mediated processes38. Furthermore, adherens junction proteins play an important role in the regulation of the actin cytoskeleton, and it was therefore hypothesized that GSK101-dependent increases in TRPV4 activity may influence both VE-cadherin proteins and actin reorganization in MTM vessel walls39-40. Consistent with this, perfusion with GSK101 followed by cell imaging indicated an apparent concentrationdependent loss of VE-cadherin immune-detection and changes to F-actin distribution (Figure 2A). Measurement of VE-cadherin fluorescence pixel intensity and distribution (% area) following perfusion with 1, 3, and 10 nM GSK101 revealed a significant reduction of VEcadherin-positive pixels across the vascular channel area when compared to control conditions (Figure 2B). Vessel perfusion with GSK101 promoted polymerization of the F-actin cortical cytoskeleton and increased stress fiber distribution proximal to the cell surface, concomitant with a loss of filamentous actin throughout the cytoplasmic regions of cells (Figure 2A). These findings in a dynamic flow 3D cell model are consistent with previous studies observed in microvascular endothelia, performed in a static 2D cell culture system23. The increase in TRPV4 activity also resulted in the formation of gaps between cells, indicating a disruption of the endothelial cell barrier. Gap measurements were determined by phalloidin-negative staining within the vessel region (relative to the total vessel area) as described previously34), and this effect was shown to be proportional to the concentration of GSK101 used (Figure 2C). The influence of TRPV4 activity on F-actin was measured by quantifying F-actin intensity and kurtosis, as a measure of F-actin distribution (larger kurtosis value indicates more peaked or discrete pixel distribution relative to a standard Gaussian distribution)41. Relative to the wide distribution of filamentous actin in the untreated control (~10.1  0.4), treatment with 1 and 3 nM GSK101 significantly increased kurtosis values (25.1  1.8; P < 0.01 and 21.1  1.7; P < 9 ACS Paragon Plus Environment


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0.05, respectively). The average F-actin fluorescence pixel intensity within the vessel area also revealed a GSK101-dependent relationship consistent with kurtosis values (Figure 2D). Treatment with 10 nM GSK101 reduced kurtosis to ~8.5  0.5 (P > 0.05) (Figure 2D), but in contrast to the filamentous distribution observed in control cells, the actin appears to be highly disorganized, leading to a significant decrease in F-actin intensity (3.59 ± 0.67-fold, P < 0.01 when compared to control conditions) (Figure 2A, D). In addition, pixel intensity line scans across individual cells (Figure 2E) showed a relatively even distribution of F-actin across cells in control conditions, enhanced actin intensity at the cell borders upon activation of TRPV4 with 1 nM or 3 nM GSK101, and the disruption of F-actin after 10 nM GSK101 treated. In summary, TRPV4-mediated Ca2+ permeation and known interactions with adherens junction proteins such as β-catenin enable TRPV4 activity and expression to control junction formation and arrangement of the actin cytoskeleton23, 30, 42-43. This is essential for barrier formation and may also play a protective role, where vascular leakiness facilitates plasma extravasation and wound healing23,

30.

A model describing the mechanisms of TRPV4-mediated control of

vascular junction formation is illustrated in Supplementary Figure S3. Together, this microfluidic system may provide an improved 3D cell culture system for understanding the role of signaling proteins such as TRPV4 in barrier integrity and vascular regulation, and provides an effective model for mimicking the leakiness of tumor vasculature, by tuning a protein that is integral to the regulation of junction formation.

Activation of TRPV4 increased vascular permeability to macromolecules. To investigate the TRPV4-mediated permeability of the vascular network, we measured the extravasation of macromolecular tracers fluorescein isothiocyanate dextran 4 kDa (FD4, ~2.8 nm in diameter) and rhodamine B isothiocyanate dextran 70 kDa (RD70; ~12 nm) from the vascular compartment into the tumor environment. These dyes are established imaging tools for


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assessing tumor edema or blood vessel barrier integrity in vivo and in microfluidic devices4447.

Following perfusion of dye solutions into the MTM for 90 min at 1.0 µL min-1, the entire tumor and vascular region of the chip was imaged, and the distribution of the dyes in vascular and tumor compartments was measured by pixel densitometry (Figure 3A). A comparison of the fluorescence distribution was measured across the tumor area (mean fluorescence intensity), to determine dye infiltration and distribution independently of fluorescence intensity. Previous co-culture MTM studies using 20kDa dextran reporters indicate that the dye can freely extravasate into the tumor area in the absence of an endothelial cell barrier 21 and have showed that significant differences in fluorescent dextran permeability can be observed within 60 min 48. In our study, 90 min of dextran uptake under non-stimulated conditions indicated saturating fluorescence of both dyes in the vascular channels, with relatively less extravasation (22.15% and 6.22% fluorescence distribution for FD4 and R70, respectively), thus confirming that a vascular endothelial cell barrier had been established (Figure 3A). Total dye accumulation (maximal fluorescence intensity) further demonstrated that extravasation of the smaller FD4 molecule from the microvascular regions into the tumor area was greater in all conditions, therefore suggesting that extravasation of the fluorescent reporters was sizedependent (Figure 3B). While optimal time points for comparing differences in dextran uptake may be achieved by time lapse studies in future investigations, 30min pre-treatment with GSK101 significantly increased dye uptake into the tumor area over the subsequent 90 min period, in a GSK101 concentration-dependent manner, providing the first indication of the relationship between TRPV4 activity and the extravasation potential of different sized macromolecules (Figure 3A, B).



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Figure 2. Activation of TRPV4 induced the formation of endothelial gaps and reorganization of F-actin. (A) Confocal images of endothelial cells stained for adherens junctions (VE-cadherin, red), Factin (phalloidin, green), and cell nuclei (Hoechst 33342, blue) after continuous perfusion of increasing concentrations of GSK101 (0, 1, 3, and 10 nM; 30 mins at 1.0 μL min-1), scale bar, 20 μm. (B) Pixel distribution above background (% area) and mean fluorescence intensity of VE-cadherin. (C) Endothelial gaps, measured by Phalloidin-negative staining within the vessel region (relative to the total vessel area). (D) Phalloidin distribution profile (relative to a standard Gaussian distribution - Kurtosis value) and mean fluorescence intensity of F-actin. Data are presented as mean ± S.E.M, n = 10 images each from 3-4 individual experiments, ns P > 0.05, * P < 0.05, ** P < 0.01, *** P < 0.001, **** P < 0.0001 versus untreated control (one-way ANOVA with Dunnett's multiple comparison test). (E) Line pixel intensity (white dashed line) and their corresponding fluorescence distribution of F-actin across a single cell (pink dashed squares in A), scale bar, 10 μm. The pixel intensity of F-actin images in 10 nM GSK101 treatment was increased by 50% to facilitate F-actin visualization.

Line pixel intensity across the horizontal axis of the tumor area was also determined to assess FD4 and RD70 fluorescence distribution and penetration into the tumor area (Figure 3C). Notably, a comparison of fluorescence intensity following MTM perfusion with GSK101 showed accumulation of FD4 with 1 nM or 3 nM GSK101, whereas significant penetration of RD70 into the tumor area was only observable after GSK101 treatment at 3 nM or 10 nM (Figure 3B). Furthermore, a 10-fold greater GSK101 concentration was required to achieve accumulation of RD70 into the tumor area comparable to levels of FD4 uptake with 1nM GSK101. Thus, the extent of vascular permeability in the MTM system could be controlled by GSK101 concentration, and 3 nM GSK101 was considered a permissive concentration to allow passage of 70kDa (12 nm) macromolecules within the assessed time period. Together, we confirmed that increasing concentrations of GSK101 led to increased gap formation between endothelial cells leading to increased vascular permeability, as evidenced by a 2-fold



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increase of FD4 and 3-fold increase of RD70 extravasation (total fluorescence uptake). Comparable macromolecule extravasation levels have been observed in previous in vivo and in vitro studies using VEGF, interleukin-2 peptide and TRPV4 agonists49-50. Using this simple, direct modulation of TRPV4 with GSK101 in a 3D microfluidic system, we can therefore tune the vascular integrity to recapitulate known TRPV4 physiological functions in vivo, especially in pulmonary edema28, 30, 51-52 and achieve vascular leakiness in a manner that is comparable with previous MTM studies using tumor cells2, 153.


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Figure 3. TRPV4 agonist-mediated changes in vascular permeability. (A) Confocal imaging of dye distribution in vessels and tumor areas, by continuous perfusion of FITC-dextran (4 kDa, green channel) and rhodamine B isothiocyanate dextran (70 kDa, red channel) for 90 mins at 1.0 µL min-1. MTMs were pre-treated with increasing concentrations of GSK101 (0, 1, 3, and 10 nM; 30 mins at 1.0 µL min-1), as indicated. (B) Pixel distribution above background (% area) and mean fluorescence intensity of FD4 and RD70 dextran dyes across the whole tumor area (area indicated by yellow dashed line). (C) Line pixel intensity across the tumor compartment (white dashed line) of FD4 (FITC-dextran) and RD70 (Rhodamine-dextran) to determine the distribution of dextran across the tumor area with increasing concentrations of GSK101. Data are presented as mean ± S.E.M, n = 3-4 individual experiments. Statistical significance from untreated control was determined by one-way ANOVA followed by Dunnett’s multiple comparisons, ns P > 0.05, ** P < 0.01, *** P < 0.001. Statistical significance between percentage area above background of FD4 and RD70 was assessed by one-way ANOVA with Tukey’s pairwise comparisons post-hoc test, # P < 0.05, # # P < 0.01. Scale bar = 100 µm.

Assessing extravasation of nanoparticles with distinct sizes and surface chemistries in an induced edema MTM model. To demonstrate the use of a MTM to rapidly screen anticancer nanocarriers, we next used fluorescence confocal microscopy and quantitative flow cytometry to track the extravasation of NPs of varying size and surface chemistry from vascular channels into tumor area and measure association with endothelial or tumor cells. Cyanine 5 (Cy5)labeled diblock copolymers of increasing size were synthesized with a PEG brush corona using the PISA method. A detailed characterization of particle synthesis and characterization was described previously54. A single macromolecular transfer agent (macro-CTA) with a carboxylic acid end group and different amounts of a hydrophobic monomer (i.e. polystyrene) were used to produce nanoparticles with diameters of 40 nm, 70 nm and 130 nm (Figure 4A). Characterization of the nanoparticle physicochemical properties (Supplementary Table S1, Figure S4) show that all nanoparticles exhibited the same spherical shape and comparable size by dynamic light scattering (DLS) and transmission electron microscopy (TEM), despite the incorporation of different macro-CTA end groups. This is consistent with our previous findings, suggesting that the macro-CTA component has minimal contribution to the overall size relative to the hydrophobic polystyrene core54. The 70 nm “medium” (M1) sized particles were also 15 ACS Paragon Plus Environment


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synthesized with a tertiary amine end group to assess the influence of surface chemistry on uptake (M1-COOH vs M2-TA; Figure 4B). A comparison of different sized carboxylic acidsurface NPs by microscopy indicated that smallest 40 nm NPs had the greatest potential for penetration into the tumor area, as measured by fluorescence pixel intensity in vascular and tumor areas (Figure 4C). Cell association with Cy5-labeled NPs determined by flow cytometry, represented by normalized median fluorescence intensity (MFI) of Cy5-positive cells, supported the imaging data, showing that 40 nm NPs (S1-COOH) had the greatest association with endothelial and tumor cells (Figure 4D and S6). This suggests that 40 nm is a permissive size that can passively move from vascular channels and into the tumor area under dynamic flow conditions. NP uptake in the absence of edema-inducing agents has previously been reported and is potentially mediated by tumor-dependent secretion of permeability-inducing molecules such as vascular endothelial growth factor from the adjacent, highly malignant breast cancer cells7, 21, 55-57. To determine the influence of surface chemistry on NP transport across endothelial barriers and accumulation into tumors in our MTM system, we compared particle distribution and cell association of 70 nm diameter NPs synthesized with carboxylic acid (M1-COOH) or tertiary amine (M2-TA) surface groups (Figure 4 and S5). Quantitative assessment by flow cytometry revealed a significantly greater association of M2-TA NPs with HUVECs. In contrast, these NPs show comparable redistribution into the tumor area and association (adherence or internalization) to tumor cells (Figure 4C, D and Supplementary Figure S6). This indicates a greater potential for tertiary amine-functionalized NPs to associate with endothelia at the vascular interface relative the 70 nm NP with a carboxylic acid on the corona (Figure 4D and S6). Together, our findings confirm the importance of size and surface chemistry for efficient NP tumor delivery and in any event where NP-cell interactions may lead to adverse effects in vitro or in vivo, these data suggest that carboxylic acid-surface NPs have a greater


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potential for delivering therapeutic agents to tumors while avoiding unwanted vascular interactions. This is consistent with our previous studies using a synthetic microvascular network to compare carboxylic acid, polyethylene glycol, methyl ester, and tertiary aminefunctionalized NPs54, and related studies using 3D tumor models, where limited tumor penetration of amine-functionalized NPs has been observed58-59.



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Figure 4. Accumulation of particles with different sizes and surface chemistries in the tumor compartment. (A) Chemical structure of diblock copolymer particles with carboxylic acid (R1) and tertiary amine (R2) end groups. (B) Schematic of tested NPs, 40, 70, and 130 nm in diameter, and 70 nm-sized particles with two different surface groups, carboxylic acid (M1-COOH) and tertiary amine (M2-TA). NPs were perfused into MTMs for 4 h followed by fixing for cell imaging, or cell dissociation for flow cytometry. (C) Fluorescence distribution of Cy5 NPs in vasculature (indicated by red dashed line) and tumor compartment (indicated by yellow dashed line), scale bars = 100 μm. Color scale, normalized fluorescence intensity. (D) Flow cytometry data showing median fluorescence intensity (MFI) relative to propodeum iodide fluorescence, in HUVECs and MDA-MB-231 cells. Data are presented as mean ± S.E.M, n = 3-4 individual experiments, * P < 0.05, ** P < 0.01, *** P < 0.001, ns P > 0.05, paired t-test.

Based on our initial characterization of endothelial TRPV4 in this microfluidic device, we subsequently measured NP extravasation and cell association in the presence of the TRPV4 agonist GSK101, as a rapid in vitro approach for assessing the EPR potential of different sized NPs when using a pharmacologically-induced edema model. Perfusion with 3 nM GSK101, was chosen due to its ability to reduce vascular barrier integrity (Figure 3), with minimal impact on cell toxicity with sustained time periods exposure (Figure S7)23. Representative images of 130 nm NPs show limited extravasation of NPs from the vascular network in control conditions, and increased punctate Cy5 fluorescence following 3 nM GSK101 treatment (30 min treatment with flow, prior to 4 h NP perfusion at 1.0 µL min-1). While NPs remained associated with endothelial cells in vascular areas under control conditions, Cy5-labelled NPs escaped the 3D vasculature with GSK101 treatment, as shown by diffusion of NPs into the porous interface between vessels and the tumor area (Figure 5A). Pixel densitometry analyses within the porous interface region showed a significant increase in Cy5 fluorescence and extravasation for all three diameter NPs, relative to the control treatment (Figure 5B). Further quantification by flow cytometry also indicated that GSK101-induced edema significantly increased extravasation of all three NPs into the tumor area, as determined by a significant increase in MDA-MB-231 cell association, while having no effect on HUVEC association (Figure 5C). Importantly, this 18 ACS Paragon Plus Environment

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demonstrates that the physical barrier present in the 2 μm porous interface does not prevent movement of NPs from the vascular channels into the tumor area. To further demonstrate the use of our MTM to screen potential anticancer nanocarriers of different sizes, we also perfused endothelial cells with increasing concentrations of GSK101 in the presence of a range of commercially-sourced polystyrene spheres with increasing diameters (20, 50, 200, and 2000 nm) and assessed their uptake using fluorescence confocal microscopy (Figure S4). While this study was not quantitative in nature, it demonstrated additional advantages for rapid, simultaneous uptake of multiple nanoparticle species in control or leaky vascular channels. Consistent with previous findings describing a therapeutic range for passive penetration of spherical nanoparticle systems between 10-100 nm60, extravasation of 20 nm spheres, and to lesser extent 50 nm spheres, was observable under control conditions. Increasing concentrations of GSK101 also increased nanoparticle extravasation of all particles and indicated size-dependent limitations to paracellular transport. For example, tumor uptake of 50 nm, 200 nm and 2 μm spheres were not observed in the tumor area under control conditions and only 50 nm spheres participated in extravasation with modest stimulation of TRPV4 activity using 1 nM GSK101. With the observed TRPV4-mediated regulation of endothelial junctions, this suggests a causative relationship between TRPV4 activity and sizedependent paracellular transport of particles. Maximal stimulation by perfusion with 10nM GSK101 only failed to promote tumour uptake of 2 μm spheres, presumably due to the restrictive size of porous barrier, which further supports the likelihood that the porous interface is not a bottleneck for particles smaller than 200 nm in diameter (Figure S4). It is noted that the breast cancer tumor cells may express the TRPV4 ion channel40, 61-62. Further studies will therefore also benefit from assessing multiple tumor cell types to confirm if the EPR effect is caused entirely by changes to endothelial permeability, or due to enhanced TRPV4-mediated association with tumor cells.



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Figure 5: GSK101 enhanced nanoparticle extravasation and accumulation into tumors. (A) Representative images showing carboxylic 130-nm particles labelled with Cy5 extravasated from the endothelial cell lining (indicated by pink dashed line) after treatment of 0 and 3 nM GSK101. HUVECs were stained with phalloidin (F-actin) and Hoechst 33342 (nucleus). (B) NP extravasation as measured percentage of fluorescence intensity of carboxylic-decorated NPs with three different sizes in the porous interface (between vascular and tumor regions) after flowing for 4 h at 1.0 μL min-1 in the MTM treated with 3 nM GSK101 (30 min, 1.0 μL min-1) over control conditions. (C) Cellular association (HUVEC and MDA-MB-231) of 40, 70 and 130nm diameter NPs measured by flow after the treatment of 3 nM GSK101 (relative to untreated control). Data are presented as mean ± S.E.M, n = 3-4 individual experiments. Statistical significance from the untreated control was determined by a paired t-test, ns P > 0.05, * P < 0.05, *** P < 0.001. One-way ANOVA with Tukey’s pairwise comparisons post-hoc test was used to assess statistical significance between intermediate- and small-sized NPs (M1-COOH


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and S1-COOH; # P < 0.05, # # P < 0.01); and large- and small-sized NPs (L1-COOH and S1-COOH; ^ P < 0.05, ^ ^ P < 0.01). Scale bar = 20 μm.

This study has exploited essential roles for TRPV4 in endothelial barrier regulation to provide insights into NP-cell interactions under physiological or edema-like conditions. TRPV4 knockdown or functional knockout is shown to reduce barrier integrity, and protein interaction studies indicating that TRPV4 can form complexes with junction and cytoskeletal proteins30,

63-65.

Hence, the endogenous presence of TRPV4 is integral for junctions to be

established, and when expressed in cells such as HUVEC and other endothelia, it can be readily modulated by selective agonists or antagonists (small molecules and lipids) and mechanical forces to tune barrier integrity. This provides a reproducible system to expedite studies optimizing nanocarriers for cancer therapy. However, several caveats must also be acknowledged. For example, while TRPV4-mediated changes in cell proliferation, migration, and angiogenesis have been observed in multiple endothelial cell types, HUVECs may not entirely recapitulate all known roles for TRPV4 in tumor vasculature24, 28, 35, 51, 53, 66. Evidence suggests TRPV4 expression levels are higher in TECs relative to normal ECs 35and

therefore may lead to greater NP extravasation if testing TECs in the MTM. Future

studies will also benefit from in vivo tumor uptake models using the same NPs tested here and a direct comparison of GSK101 perfusion with endogenous edema mediators such as interleukins, thrombin, or VEGF, or tumor supernatants that are likely to contain lipid-based TRPV4 activators such as arachidonic acid should also be tested, to confirm if the use of a selective TRPV4 agonist recapitulates the cytoskeletal arrangement and angiogenesis previously observed in endothelia21, 35, 53, 67. It should also be noted that exposing cells and particles to variable flow rates to simulate the heterogeneity of shear forces expected in vivo, can only be perceived as a theoretical advantage until rigorous analyses and comparison are made between branched and straight channel devices. Indeed, further simplification may also



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be achieved through use of straight channel systems for rapidly identifying nanoparticle properties that are desirable for avoiding vascular cellular interactions and enhanced tumor penetration when assessing TRPV4 as an edema mediator. Together, stimulation of TRPV4 in a microfluidic device seeded with endothelial cells offers a simple, animal-free, biologically relevant MTM that is scalable and relatively cost-effective, with potential to rapidly screen extravasation and tumor accumulation of nanocarriers. These data demonstrate that stimulation of TRPV4 activity can readily modulate the endothelial cell barrier to assess NP extravasation in leaky vessels without loss of cell viability, to offer a highly tunable system for modelling cancers with variable levels of vascular leakiness7. Multiple nanoparticles can be simultaneously assessed and the combination of imaging with flow cytometry provides both qualitative and quantitative approaches for measuring nanoparticle uptake, for rapid assessment of particle properties, such as size and surface chemistry, to optimize their EPR potential.

CONCLUSIONS In this study, a tumor-on-a-chip microfluidic system incorporating a tumor area bordered by branched endothelial vasculature was used as a model to screen nanoparticles and macromolecules with unique physicochemical properties, in order to rapidly identify those properties that may be beneficial for extravasation, tumor penetration and cancer therapy. Using this system, we first confirmed that NP size is a critical factor for passage across endothelial barriers and tumor accumulation, by comparing different diameter carboxylic acidterminated particles to reveal a higher degree of extravasation and tumor penetration with 40 nm particle, relative to the larger counterparts (70 and 130 nm). Furthermore, while a comparison of 70 nm-sized NPs bearing carboxylic acid and tertiary amine surface groups revealed no significant difference in movement of particles into the tumor area, surface chemistry significantly influenced endothelial cell association, thus indicating that the 22 ACS Paragon Plus Environment

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carboxylic acid surface may be favored therapeutically for penetrating the vascular barrier and avoiding unwanted endothelial interactions. The MTM system provided a valuable in vitro co-culture cell system with a physiologically relevant 3D cellular architecture under flow conditions, that may be superior to 2D culture systems for exploring the role of TRPV4 in endothelial barrier formation. Studies in model HUVEC cells showed that the activation of TRPV4 by the specific agonist GSK101 promoted the rearrangement of F-actin cytoskeleton most likely through Ca2+-dependent processes35, leading to reduced detection of VE-cadherin and subsequent disruption of intercellular junctions between endothelial cells. Further studies in this system may also provide new insights into the role of TRPV4 activity in edema, and supports the hypothesis that TRPV4 is a potential therapeutic target for regulating nanoparticle accumulation and retention in tumors. The signaling processes of multiple pro-edema targets such as bradykinin and protease-activated receptors can also converge on TRPV4 to regulate its activity68-69, thus indicating the potential for additional unappreciated roles for TRPV4 in tumor vascular permeability that could be investigated further in MTMs. The TRPV4 agonist induced formation of intercellular gaps allowing different levels of leakiness in a concentration-dependent manner and this was confirmed using macromolecular dyes, commercial spheres and synthesized polystyrene-based Cy5-labeled NPs. With the possibility of real-time visualization and quantification of the endothelial permeability and tumor accumulation, the MTM system with “induced edema” is a useful in vitro model to rapidly screen potential anticancer nanocarriers intended to exploit the EPR effect. While the present model could not fully represent the complexity of biological systems (e.g., lack of plasma proteins or circulating blood cells) that are known to affect NP properties and nano-bio interactions70-71, the ease and expected consistency of setting up this microfluidic system has broad utility in multiple cell barrier systems where TRPV4 is expressed (e.g.



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vascularized tumor, colon epithelia & blood brain barrier). In particular, this is proposed to be valuable as a pre-screening tool for materials-focused studies that require rapid characterization of nanoparticle interactions and efficacy, before advancing studies into more biological complex systems. Microfluidic studies using linear channels have also shown that use of coculture devices to screen NPs and determine optimal properties such as size, shape and surface charge may be translated to in vivo tumor models27 and may also be further exploited via pharmacological regulation of edema-inducing signaling targets such as TRPV4.

EXPERIMENTAL SECTION Reagents. All reagents and consumables were purchased from Sigma Aldrich, unless stated otherwise. Cell culture in microfluidic tumor models. Prior to seeding cells in the microfluidic tumor chip (SynVivo, 105007-STu), channels and tumor area were coated with Matrigel® (In Vitro Technologies) in chilled serum-free endothelial cell basal media (EBM; 1:10 dilution, Lonza, Australia) and incubated for 1 h at 4°C followed by 1 h at 37 °C in 5% CO2 and washed with serum-free EGM. Breast cancer cell lines MDA-MB-231 (ATCC) were maintained as monolayers in filter cap flasks in RPMI-1640 medium supplemented with 10% (v/v) of fetal bovine serum and 1% penicillin/ streptomycin (Life Technologies) at 37 °C, 5% CO2. Using a cell suspension of 1 × 107 cells mL-1, cells were slowly pipetted into the tumor compartment of the chip until they filled ~80-90% of the area. The cells were then incubated for 1 h (37 °C in 5% CO2) to allow the cells to attach before perfusion with fresh media (0.1 μL min-1). Cells were maintained for 24 h before seeding Human Umbilical Vein Endothelial Cells (HUVECs; Lonza) into the vascular network. HUVECs were maintained in EBM supplemented with Endothelial Cell Growth Media SingleQuots (Lonza) in a humidified incubator at 37 °C, 5 % CO2. Cells were harvested and diluted 2.5 × 107 cells mL-1in EBM, then slowly injected by 24 ACS Paragon Plus Environment

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hand into the vascular networks using a sterile syringe until covering 80-90% of the vessel surface, then incubated for 2 h at 37 °C, 5 % CO2. EGM was then perfused through the MTM at 0.1 μL min-1 for a minimum of 22 h using a syringe pump (Harvard Apparatus, Holliston MA) and cells were grown to confluence to establish a vascular cell barrier. For initial imaging of MDA-MB-231 GFP fluorescence, cells were fixed by perfusion with 4% paraformaldehyde (PFA) in PBS, perfused into the MTM for 15 min at room temperature and stained with 0.3 nM Hoechst 33342. Immunofluorescent staining. Immunostaining was performed using antibodies selective for endothelial proteins VE-cadherin (CD144) and PECAM1 (CD31). Briefly, cells were fixed by perfusion with 4% PFA in PBS (15 min, room temperature), then treated with 0.1% Triton X100 in PBS (10 min, room temperature) and blocked by perfusion with 1% bovine serum albumin (BSA), 0.3 M glycine (Abcam) in 1% PBS-Tween 20 (Abcam) and incubation for 1 h (room temperature), followed by incubation in blocking solution with rabbit polyclonal antibodies against VE-cadherin or CD31 (Abcam) (1:400) overnight at 4 °C, secondary goat anti-rabbit IgG Alexa Fluor 594 (1:1000, Abcam) and Phalloidin-iFluor 488 (1:1000, Abcam) for 2 h at room temperature, then incubated with 0.3 nM Hoechst 33342 for 15 min at room temperature. After each step, the cells were washed by perfusion at 1 μL min-1, with PBS for 5 min. Fluorescence Microscopy. Fluorescent and bright-field images were acquired on a Nikon Eclipse TiE inverted microscope equipped with a A1R Resonant Scanning Confocal system, using 10× 0.30 NA Plan Fluor or 20× 0.45 NA SPlan Fluor ELWD objective with 4× magnification, and excitation/emission 405/ 45025 (blue channel); 488/52525 (green channel), 561/59525 (red channel) for immunofluorescent staining, or Cy5 fluorescence using 640/663-738 for nanoparticle uptake. All images were captured at 1024×1024 pixels resolution using a pixel dwell time of 2.4 µs/pixel. 25 ACS Paragon Plus Environment


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Image Processing. Images were processed initially using NIS Elements software (version 4.30, Nikon Instruments) followed by FIJI software 72. Percentage area of VE-cadherin or F-actin was calculated by measuring VE-cadherin or F-actin-positive pixels relative to the total area, for vascular channels randomly imaged from minimum of 3 individual experiments. Percentage of endothelial gaps was quantified by measuring phalloidin-negative pixels (no fluorescence) across a region of interest, divided by total area of imaged vessels. Mean fluorescent intensity of VE-cadherin and F-actin was determined in FIJI by drawing a region of interest around the entire tumor area. The analysis was performed using FIJI software with at least 10 images with about 100 cells taken from three different chips for each condition. Data are presented as mean ± S.E.M. Nanoparticle and Dextran reporter dye extravasation and accumulation into tumors. To induce edema effects, the MTM was perfused with TRPV4 agonist GSK1016790A (GSK101) at 1, 3, and 10 nM in EGM at 1.0 μL min-1 for 30 min at 37 °C, 5% CO2. For dextran uptake, a 25 µM solution of fluorescein isothiocyanate dextran 4 kDa or rhodamine B isothiocyanate dextran 70 kDa (Sigma-Aldrich) in EGM was perfused at 1.0 µL min-1 for 90 min. To capture large regions of the chips, live cells were imaged with a 10× 0.30 NA Plan Fluor objective. Percent area of dyes was quantified as number of pixels above background (FITC or Rhodamine) divided by total pixels within the chip tumor area. Mean pixel intensity of dextran in whole tumor regions and their line pixel intensity across long axis of tumor compartment was determined using FIJI software (n = 3 or more individual experiments). For uptake of three different sized Cy5-labeled particles (~40, ~70, and ~130 nm), a 240 μL EGM suspension particles in equivalent numbers (~ 1 × 1011 NPs mL-1) were perfused through the MTM system at 1.0 μL min-1 using a syringe pump, with and without prior perfusion with 3 nM GSK101 (1.0 μL min-1, 30 min). After 4 h, 1 mL of chilled HBSS was used to wash out unassociated particles, followed by 0.3 mM Hoechst 33342 labelling (15 min at 5.0 μL min-1)


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15 min incubation at 4 °C before fixation and staining with Hoechst 33342 and Phalloidin. To account for differences in the stoichiometry of Cy5 fluorophore incorporation into different sized particles, fluorescence intensity was normalized according to spectrophotometer measurements of equivalent particle numbers (Figure S3). Nanoparticles synthesis. Carboxylic-terminated particles of three different sizes (~40, ~70, and ~130 nm), and intermediate-sized particles with tertiary amine end groups were synthesized using polymerization-induced self-assembly (PISA) technique as described previously.41 Briefly, the corona of the diblock copolymer particles was synthesized by reversible addition-fragmentation chain transfer (RAFT) solution polymerization of Nhydroxyethyl acrylamide (HEAA, 97%, Sigma-Aldrich) and poly(ethylene glycol) methyl ether

acrylate

(PEGA,

average

Mn

=

480;

Sigma-Aldrich)

with

4-cyano-4-

(ethylthiocarbonothioylthio) pentanoic acid (ECT) or amine-modified ECT in DMSO for 4 h at 70 °C to obtain macromolecular chain transfer agent (macro-CTAs). The core of the particles was formed by chain extension of macro-CTA with styrene using RAFT-mediated emulsion polymerization in water at 80 °C. After 4 h polymerization, an aliquot of Cy5-maleimide in styrene was added into the reaction using a gas tight syringe. The reaction was continued for a further 2 h to allow incorporation of Cy5 into the core of the micelles. Nanoparticle characterization. Dynamic Light Scattering (DLS): A solution of nanoparticles (50 μg mL-1 in water and EGM) was measured by a Malvern Zetasizer Nano Series running DTS software and operating a 4 mW He–Ne laser at 633 nm. Data measurement and analysis was performed at an angle of 173° and a temperature of 25 °C, to report the number average hydrodynamic diameter and polydispersity index. Fluorescence measurement: To assess differences in fluorophore incorporation for different sized nanoparticles, fluorescence spectra of a solution of nanoparticles with equivalent numbers was obtained using a fluorescence spectrophotometer (Shimadzu RF-501 PC) at an excitation wavelength of 635 nm, and slit



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widths of 10 nm for both excitation and emission of Cy5. Transmission electron microscopy (TEM): An aliquot (5 µL) of 0.1 wt% NP solution in MiliQ water was deposited on a Formvarcoated copper grid (GSCu100F-50, Proscitech) and left to dry overnight in air and at room temperature. TEM imaging was performed using a Tecnai F20 transmission electron microscope at an accelerating voltage of 200 kV at ambient temperature, as described previously.41 Flow cytometry. Cell were dissociated with chilled 0.025% trypsin/EDTA and incubated at 4 °C for 5 min then HUVECs and MDA-MB-231 were harvested separately out of the top of the gel basement via two different ports. Cells were centrifuged (300 g, 5 min, 4 °C) and resuspended in 300 μL of 1 mM propidium iodide (PI) in HBSS before being measured Cy5 fluorescence (ex/em: 633/66020) in PI-negative cells (488/670 long pass) by flow cytometry using a FACS Canto II (BD Biosciences, San Jose, CA). All experiments were performed in triplicate, from 3 or more experiments.


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ASSOCIATED CONTENT Supporting Information. The Supporting Information is available free of charge on the ACS Publications website at DOI: Cytotoxicity of GSK101 on the cells; formation of adherens junctions in HUVECs; model for regulation of VE-cadherin and F-actin by TRPV4 activation; physicochemical properties of nanoparticles, flow cytometry raw data; rapid screening of size-dependent particle accumulation AUTHOR INFORMATION Corresponding Authors *Email: [email protected] & [email protected]

Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Conflict of interest The authors declare no competing financial interest. ACKNOWLEDGMENTS We would like to thank Dr. Angus Johnston for support with data analysis and Mr. Daniel Brundel for providing materials for cell maintenance and imaging. This work was supported by the Australian Research Council (ARC) Centre of Excellence in Convergent Bio-Nano Science and Technology (Project No. CE140100036), the Vietnamese Government and the Monash University Faculty of Pharmacy and Pharmaceutical Sciences (to M.N.V.), the Australian Government Research Training Program scholarship (to S.Y.K.), ARC DECRA 29 ACS Paragon Plus Environment


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Fellowship to N.P.T. (DE180100076), Takeda Pharmaceuticals Inc. (to N.V) and award of an ARC Australian Laureate Fellowship to T.P.D (FL140100052). REFERENCES 1. Spill, F.; Reynolds, D. S.; Kamm, R. D.; Zaman, M. H., Impact of the Physical Microenvironment on Tumor Progression and Metastasis. Curr. Opin. Biotechnol. 2016, 40, 41-48. 2. Maeda, H.; Wu, J.; Sawa, T.; Matsumura, Y.; Hori, K., Tumor Vascular Permeability and the EPR Effect in Macromolecular Therapeutics: A Review. J. Controlled Release 2000, 65, 271-284. 3. Matsumura, Y.; Maeda, H., A New Concept for Macromolecular Therapeutics in Cancer Chemotherapy: Mechanism of Tumoritropic Accumulation of Proteins and the Antitumor Agents Smancs. Cancer research 1986, 46, 6387- 6392. 4. Wilhelm, S.; Tavares, A. J.; Chan, W. C. W., Reply to "Evaluation of Nanomedicines: Stick to the Basics". Nat. Rev. Mat. 2016, 1, 16074 5. Wilhelm, S.; Tavares, A. J.; Dai, Q.; Ohta, S.; Audet, J.; Dvorak, H. F.; Chan, W. C. W., Analysis of Nanoparticle Delivery to Tumours. Nat. Rev. Mat. 2016, 1, 16014. 6. Prabhakar, U.; Maeda, H.; Jain, R. K.; Sevick-Muraca, E. M.; Zamboni, W.; Farokhzad, O. C.; Barry, S. T.; Gabizon, A.; Grodzinski, P.; Blakey, D. C., Challenges and Key Considerations of the Enhanced Permeability and Retention Effect for Nanomedicine Drug Delivery in Oncology. Cancer Research 2013, 73. 7. Hobbs, S. K.; Monsky, W. L.; Yuan, F.; Roberts, W. G.; Griffith, L.; Torchilin, V. P.; Jain, R. K., Regulation of Transport Pathways in Tumor Vessels: Role of Tumor Type and Microenvironment. Proc. Natl. Acad. Sci. U. S. A. 1998, 95, 4607-4612. 8. He, C.; Hu, Y.; Yin, L.; Tang, C.; Yin, C., Effects of Particle Size and Surface Charge on Cellular Uptake and Biodistribution of Polymeric Nanoparticles. Biomaterials 2010, 31, 3657-66. 9. Truong, N. P.; Whittaker, M. R.; Mak, C. W.; Davis, T. P., The Importance of Nanoparticle Shape in Cancer Drug Delivery. Expert Opin. Drug Delivery 2015, 12, 129-142. 10. Agarwal, P.; Wang, H.; Sun, M.; Xu, J.; Zhao, S.; Liu, Z.; Gooch, K. J.; Zhao, Y.; Lu, X.; He, X., Microfluidics Enabled Bottom-Up Engineering of 3D Vascularized Tumor for Drug Discovery. ACS Nano 2017, 11, 6691−6702. 11. Cox, M. C.; Reese, L. M.; Bickford, L. R.; Verbridge, S. S., Toward the Broad Adoption of 3D Tumor Models in the Cancer Drug Pipeline. ACS Biomater. Sci. Eng. 2015, 1, 877-894. 12. Pradhan, S.; Smith, A. M.; Garson, C. J.; Hassani, I.; Seeto, W. J.; Pant, K.; Arnold, R. D.; Prabhakarpandian, B.; Lipke, E. A., A Microvascularized Tumor-mimetic Platform for Assessing Anti-cancer Drug Efficacy. Sci. Rep. 2018, 8, 3171. 13. Breslin, S.; O'Driscoll, L., Three-Dimensional Cell Culture: The Missing Link in Drug Discovery. Drug Discovery Today 2013, 18, 240-249. 14. Kwak, B.; Ozcelikkale, A.; Shin, C. S.; Park, K.; Han, B., Simulation of Complex Transport of Nanoparticles Around a Tumor Using Tumor-Microenvironment-on-Chip. J. Controlled Release 2014, 194, 157-167. 15. Sackmann, E. K.; Fulton, A. L.; Beebe, D. J., The Present and Future Role of Microfluidics in Biomedical Research. Nature 2014, 507, 181-9.


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Microfluidic device for rapid assessment of nanoparticle extravasation and tumor uptake

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Rapid Assessment of Nanoparticle Extravasation in a Microfluidic

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