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ACS Appl Mater Interfaces
ACS Appl Mater Interfaces
am
aamick
ACS Applied Materials & Interfaces
1944-8244
1944-8252
American Chemical Society

39196896
10.1021/acsami.4c01766
Research Article
Magnetically Integrated Tumor–Vascular Interface System to Mimic Pro-angiogenic Endothelial Dysregulations for On-Chip Drug Testing
Surendran Vikram †
Safarulla Simrit †
Griffith Christian ‡
Ali Reem †
Madan Ankit §
Polacheck William ‡
https://orcid.org/0000-0001-5205-3591
Chandrasekaran Arvind *†
† Bioinspired Microengineering (BIOME) Laboratory, Department of Chemical, Biological and Bio Engineering, North Carolina A&T State University, Greensboro, North Carolina 27265, United States
‡ Joint Department of Biomedical Engineering, UNC Chapel Hill—NC State University, Chapel Hill, North Carolina 27599, United States
§ MedStar Southern Maryland Hospital Center, MedStar Georgetown Cancer Institute, Clinton, Maryland 20735, United States
* Email: achandra@ncat.edu. Phone: + 1 336 5542111.
28 08 2024
11 09 2024
16 36 4707547088
06 02 2024
29 07 2024
18 06 2024
© 2024 The Authors. Published by American Chemical Society
2024
The Authors
https://creativecommons.org/licenses/by-nc-nd/4.0/ Permits non-commercial access and re-use, provided that author attribution and integrity are maintained; but does not permit creation of adaptations or other derivative works (https://creativecommons.org/licenses/by-nc-nd/4.0/).

The tumor–vascular interface is a critical component of the tumor microenvironment that regulates all of the dynamic interactions between a growing tumor and the endothelial lining of the surrounding vasculature. In this paper, we report the design and development of a custom-engineered tumor–vascular interface system for investigating the early stage tumor-mediated pro-angiogenic dysfunctional behavior of the endothelium. Using representative endothelial cells and triple negative breast cancer cell lines, we established a biomimetic interface between a three-dimensional tumor tissue across a mature, functional endothelial barrier using a magnetically hybrid-integrated tumor–vascular interface system, wherein vasculature-like features containing a monolayer of endothelial cell culture on porous microfluidic channel surfaces were magnetically attached to tumor spheroids generated on a composite polymer-hydrogel microwell plate and embedded in a collagen matrix. Tumor-mediated endothelial microdynamics were characterized by their hallmark behavior such as loss of endothelial adherens junctions, increased cell density, proliferation, and changes in cell spreading and corroborated with endothelial YAP/TAZ nuclear translocation. We further confirm the feasibility of drug-mediated reversal of this pro-angiogenic endothelial organization through two different signaling mechanisms, namely, inhibition of the vascular endothelial growth factor pathway and the Notch signaling pathway, thereby demonstrating the utility of the tumor–vascular interface platform for rapid, early stage prediction of antiangiogenic drug efficacy. Overall, our work emphasizes the importance of our strategic engineering approach for identifying some unique, physiologically relevant aspects of the tumor–vascular interface, which are otherwise difficult to implement using standard in vitro approaches.

tumor–vascular interface
tumor endothelium
hybrid integration
magnetic attachment
angiogenesis
endothelial YAP/TAZ
VEGF inhibitor
notch signaling
National Institutes of Health 10.13039/100000002 T32HL69768 North Carolina Translational and Clinical Sciences Institute, University of North Carolina at Chapel Hill 10.13039/100011485 550KR282124 Defense Human Resources Activity 10.13039/100010210 W911NF2120265 American Heart Association 10.13039/100000968 CDA857738 National Institute of General Medical Sciences 10.13039/100000057 5SC2GM136523 document-id-old-9am4c01766
document-id-new-14am4c01766
ccc-price
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pmc1 Introduction

Targeting tumor angiogenesis through therapeutic interventions is emerging as a promising early stage treatment option for cancers and is extensively being investigated in preclinical and clinical settings.1−3 Our overall understanding of the process of tumor angiogenesis has largely been aided by the numerous in vitro models that are currently available,4−6 given the challenges of using animal models to dissect the complex underlying events surrounding the tumor growth and metastasis. However, for successful implementation of the in vitro models as surrogates to animal testing or human clinical trials, it is important to identify the critical components of a tumor-specific microenvironment and the underlying cellular level intricacies to be incorporated within these models, which would help gain a focused understanding of the overall translational biological relevance. The tumor–vascular microenvironment (TVME) is the basic functional unit of the tumor etiology,7,8 and the interface between malignant tumors and the nontransformed cells of the surrounding vasculature, also referred to as the tumor–vascular interface, is a vital subunit of the TVME, which can regulate the tumor growth, dissemination, and resistance to therapeutics at various stages of carcinogenesis.9 The emergence of key findings highlighting the major role played by the tumor–vascular interface is increasingly being discussed as potential targets for the development of early stage antiangiogenic therapeutic strategies.10,11 This gives the motivation to develop robust tumor–vascular interface models for conducting physiologically relevant in vitro studies of angiogenesis and cancer metastasis.

One of the important features of a tumor–vascular interface is the monolayer of endothelial cells (ECs) lining the vasculature.9 The endothelial lining not only acts as a barrier between tumor tissue and blood but also regulates important and dynamic tissue functions, including regulation of tumor cell intravasation, vascular dynamics, angiogenesis, and transport of fluid and soluble factors, among others.12 At the tumor–vascular interface, aggressive development of the tumor and the resultant metabolic demands mediate an overexpression of pro-angiogenic factors, which triggers aberrant vascular microdynamics,13 typically characterized by the disorganization of the endothelial lining into an immature, proliferative, and permeable phenotype (schematically shown in Figure 1A) that subsequently induces angiogenesis and promotes metastasis.14 While an increasing number of research works have identified and validated factors and signaling pathways that govern tumor-mediated vascular dysfunctions under angiogenesis conditions, the mechanisms of action and relative importance of these factors, particularly in driving the dysregulated vascular dynamics of an early stage tumor, remain largely unknown.15,16 In this paper, we investigate the onset of early stage tumor-mediated pro-angiogenic vascular endothelial dysfunction and drug-mediated reversal of this behavior, using a hybrid-engineered coculture system that can recapitulate the in vivo physiology of a tumor–vascular interface.

Figure 1 (A) Schematic representation of a tumor–vascular interface showing tumor-induced pro-angiogenic endothelial phenotypes including increased proliferation and loss of adherens junctions leading to increased vascular permeability. (B) Schematic cross-sectional conceptualization of the tumor–vascular interface using magnetic hybrid integration: tumor spheroids embedded within a collagen matrix and housed within a composite polymer-hydrogel microwell assembly are magnetically attached to an endothelialized microfluidic channel fabricated on the porous membrane of a custom-made Transwell insert and placed inside a support vessel.

To study the dynamics of an early stage tumor vasculature in vitro, it is imperative to integrate the vascular interface into existing tumor model16,17 for developing physiologically relevant representations of a growing tumor. However, designing the complexities of a tumor–vascular interface has always remained a challenge. A tumor–vascular interface model can be successful only if it is adequately designed and controlled since it involves unique organization and fabrication requirements. These include the need to move toward three-dimensional (3D) approaches,18 solid tumors,19 apical–basal polarity of the endothelial lining,20 perfusion of blood vessels,21 etc. For example, drugs targeting solid tumors must penetrate the tumor tissues, passing across the vascular endothelial lining, and this process heavily depends upon the apicobasal organization of the endothelial monolayer at the tumor–vascular interface.19 Consequently, bioengineering of a functional tumor–vascular interface has, in general, been a major challenge22 because successful vascularization of a tumor necessitates the integration of multiple tissue microenvironments including 3D tumoroids, stroma, and active vascular conduits lined with functional ECs in a dynamic and biochemically relevant coculture format. Simplistic models of the tumor-vasculature interface based on tumor-endothelial cocultures fabricated using planar, two-dimensional culture vessels may not accurately recapitulate these tissue-specific dynamics of the TVME under normal and pathological conditions. The existence of other technical issues related to reproducibility, lack of a realistic tumor microenvironment, long-term viability, difficulty of handling and implementation, and post-assay analytical limitations involving multiple cell types and behaviors could also restrict the feasibility of conducting complex assays using such models. Given that EC dynamics are highly sensitive to tumor-regulated paracrine interactions, which evolve during tumor growth, improved physiological relevance of an in vitro tumor–vascular interface necessitates establishing a functional and perfusable vascular endothelium prior to introduction of the tumor. Such a strategy to create a biomimetic coculture that enables temporal control of tumor-mediated signals would allow controlled studies of the vascular abnormalities resulting from tumor growth and, conversely, the impact of the vasculature on the tumor dynamics.

A common approach for generating spatially segregated tumor-endothelial coculture has been Transwell culture in multiwell plate formats.4,16 With ECs on the apical side of a Transwell membrane and tumor cells cultured on the basolateral side, Transwell models are capable of maintaining a coculture with controlled intercellular contact and facilitating the formation and maturation of EC–cell junctions, which are required for vascular homeostasis and establishment of apical–basal polarity and proper deposition of the vascular basement membrane.23 Transwells provide access to both the apical and basal compartments for medium sampling or drug application, facilitating transendothelial electrical resistance (TEER) measurements,24 a gold standard for nondestructive quantification of endothelial barrier integrity. Furthermore, the optical transparency of Transwell membranes enable visualization of the cell dynamics over the course of the experiments.25 However, major limitations to commercial Transwell systems such as the lack of design customizability required for 3D tumor cultures26 and the lack of pressure and flow control for perfusion studies27 preclude their use in the development of physiologically relevant tumor–vascular interface models. Aided by advancements in tissue engineering and microengineering technologies,28 a number of approaches have been adopted for the bioengineering of tumor-vasculature-on-chips (TVoCs).29 TVoCs are miniaturized coculture platforms wherein 3D tumoroid models are grown within synthetic or reconstituted extracellular matrix and integrated with vascular networks to establish tumor–vascular interfaces.30−33 The incorporation of microfluidics into the TvoCs34−36 facilitates perfusion of the vascular networks and associated paracrine signaling between endothelial and tumor cells. Thus, TVoCs have found a wide range of applications, including evaluation of tumor-mediated phenomena such as angiogenesis,37 cell migration, drug delivery,38 and drug-screening.39 However, TVoC models often require complex fabrication strategies and result in devices that do not allow end-point cell separation or nondestructive retrieval of cells or tissues, which further impedes the ability to conduct TEER measurements.40,41 While attempts have been made to fabricate microfluidics on Transwell membranes,27 size limitations and fabrication complexity limit throughput and pose technical challenges for routine tasks including replenishing nutrients and introducing cells and tissues with spatiotemporal resolution. These drawbacks emphasize the need for more user-friendly, easy-to-prototype, high-throughput, scalable, and physiologically relevant tumor–vascular interface models that could still retain compatibility with existing analytical tools or imaging systems and thus allow for reliable usage over a wide range of research applications.

Here, we leverage the advantages of Transwell cultures and TVoC models in the development of a versatile, magnetically integrated tumor–vascular interface system (Figure 1B). This hybrid-engineered coculture design consists of vascular ECs cultured within a microfluidic channel created on top of a custom-designed Transwell membrane, which is magnetically aligned and attached to a polymer-hydrogel composite microplate assembly that consists of an array of tumor spheroids embedded within collagen matrix. In this arrangement, ECs are cultured in proximity to 3D tumor spheroids and maintain apical–basal polarity, recapitulating the in vivo physiology of a tumor–vascular interface.42 The use of a custom magnetic system provides on-demand attachment and detachment of the coculture ensembles to enable temporal control over coculture initiation and subsequent segregation of cells and tissue for downstream analysis. Using human umbilical vein ECs (HUVECs) and triple-negative breast cancer cells (MDA-MB-231), we implemented the tumor–vascular interface to investigate the modalities of early stage tumor-mediated pro-angiogenic vascular dysfunctions. We found that the presence of tumor cells significantly affects the proliferation, spreading dynamics, and cell–cell adherens junction integrity of ECs, along with nuclear translocation of the endothelial yes-associated protein (YAP and the transcriptional coactivator with PDZ-binding motif (TAZ). We further confirm the feasibility of drug-mediated reversal of these tumor-mediated pro-angiogenic endothelial organization through two different signaling mechanisms, thereby demonstrating the utility of the magnetically integrated tumor–vascular interface platform for rapid, early stage prediction of antiangiogenic drug efficacy.

2 Materials and Methods

2.1 Design of the Tumor–Vascular Interface

The principal components of the magnetically integrated tumor–vascular interface chip (henceforth called as M-TuVIC) are the tumor spheroid assembly, the vascular endothelial layer, and the “stromal” collagen layer in between the endothelium and the tumor (Figure 2A). The vascular component consists of an endothelialized microfluidic channel surface created on a hydrophilic porous membrane attached to a custom-designed 3D printed Transwell insert. Using ring magnets, this assembly is integrated with a tumor spheroid array generated within hydrogel microwells and embedded within a collagen matrix43,44 and placed inside a customizable support vessel, similar to a Transwell insert assembly (as schematically shown in Figure 1B).

Figure 2 (A) Implementation of M-TuVIC: using ring magnets, microfluidic channel printed on a porous membrane of a custom-made Transwell insert is integrated with a composite polymer (PDMS)-hydrogel microwell plate housing the spheroids, closed with a monolithically built-in hemispherical seal around the microwell plate providing a leak-proof assembly. (B) Fabrication and assembly of the M-TuVIC: (i) microfluidic channel on the porous membrane is fabricated by printing a sacrificial pluronic layer on the porous membrane, followed by creation of the channel enclosure using UV-curable glue. The setup is attached to the custom-made Transwell insert using a temporary magnetic assembly, after which the pluronic layer is washed away to create the channels. ECs are thereafter cultured in the channel until formation of a mature barrier (∼7 days). (ii) Process of fabricating the tumor spheroid assembly starts with 3D printing custom molds for replica molding of the hydrogel microwells,63 which is grafted onto a PDMS base (Supporting Information Figure S1). For visibility, the PDMS base can be color-coded. Tumor spheroids are generated inside the microwells by self-aggregation of the tumor cells prior to hybrid integration. (iii) Endothelial assembly and the tumor spheroid assembly are integrated by aligning a ring magnet under the PDMS disk over a flat surface and positioning the Transwell assembly (containing the microfluidic channel and a ring magnet placed inside) from the top.

The space between the spheroids and the endothelialized channel is filled with collagen type I,43 which forms the stromal region. The hydrogel microwells are grafted onto a perforated polymer disk with monolithically fabricated hemispherical gaskets (Supporting Information Figure S1). These disks enable handling the hydrogel during the assays, and the built-in gaskets provide a leak-proof sandwich of the collagen layer between the microwells and the Transwell membrane under the magnetic attachment. The geometric distance between the ECs and the spheroids is thus governed by the thickness of the hemispherical gasket seals. The perforations on the polymer disk enable the transport of culture media between the support vessel and the collagen layer, necessary to maintain the tumor spheroid culture. For all of the experiments in this work, 6-well plates were used as the support vessels, and the Transwell inserts for the M-TuVIC were custom designed so as to fit into the 6-well plates within the working distance available to accommodate the magnetic attachment of the microwell plate assembly from the bottom of these inserts. Overall, the design of the M-TuVIC allows for the culture of the ECs and the tumor spheroids separately before attaching these components to enable the tumor–vascular interface.

2.2 Fabrication of the M-TuVIC

2.2.1 Rapid Prototyping of Porous Microfluidic Channels

Microfluidic channels were fabricated on a porous membrane by bioprinting a sacrificial channel layer that upon creation of a surrounding enclosure layer was subsequently dissolved to leave behind conduits that enable fluid flow over a permeable substrate (Figure 2Bi). A commercial 3D bioprinter (Bio X, CellInk, VA, USA) was used to print a sacrificial layer of pluronic gel (IK50000, CellInk, USA) in the desired contour of the microfluidic channel. Initial calibration of the 3D bioprinter was conducted to optimize the ink extrusion parameters and to achieve the desired channel dimensions. After printing the pluronic layer, microfluidic tubes (EW-06419–01, Cole Parmer, USA) were stably positioned on the channel ends for creating the channel inlet and outlet. UV-curable polymer NOA63 (Norland Inc., USA) was then cast on top of the pluronic gel, surrounding and enclosing the channel features. The setup was then exposed to 365 nm UV light for 5 min to completely polymerize and harden the NOA63 layer. The assembly was then cooled to 4 °C for 15 min, after which the sacrificial pluronic gel was washed off by flushing the inlet and outlet tubes with cold phosphate-buffered saline (PBS). Unless otherwise specified, straight microfluidic channels with 350 μm width and 350 μm height were used in our assays. For the M-TuVIC configuration used in endothelial proliferation assays, the width of the endothelial channel was at least 5 times greater than the diameter of the tumoroids.

To accommodate the “chopstick” TEER probes (4 mm wide × 1 mm thick, World Precision Instruments, USA) inside the microchannel for conducting TEER measurements, open microfluidic channels with 1.5 mm width were fabricated by applying minor changes to the procedure described above. Since the ECs are grown on the surface of the channel, the operational principles between the two devices are the same, and the TEER-compatible device can be replaced with the standard microfluidic design for any perfusion-based assays. For the open channel fabrication (Supporting Information Figure S2), the thickness of the sacrificial pluronic layer was increased, and the UV-curable glue was precisely bioprinted around the pluronic channel without creating any enclosure on top of the pluronic gel. Cold PBS added on top of the assembly dissolved the pluronic ink, creating an open conduit on top of the porous membrane that acts as a defined culture space for that ECs that could also facilitate TEER analysis.

2.2.2 Fabrication of Composite Microwell Plate Assembly

To fabricate the composite microwell plate assembly, polyacrylamide (PAAm) was used to generate and culture tumor spheroids, and the supporting perforated disks were fabricated from polydimethylsiloxane (PDMS). During the process of replica molding of the microwells, the PAAm gel was grafted45 onto the preformed PDMS disks to create the composite assembly.

To fabricate monolithic PDMS disks with built-in hemispherical seals, a plastic mold (with inner diameter of 10 mm) consisting of circular tapered groove (13 mm diameter, 0.5 mm tall, 1 mm wide) was 3D printed (Formlabs 3, Figure 2Bii). The molds were spray-coated with ease-release 200 (Mann Release Technologies) to facilitate easy removal of PDMS from 3D printed parts. Sylgard 184 PDMS (Dow Corning) was mixed in a volumetric 10:1 base to cross-linker ratio, degassed, and poured onto the plastic mold. Since PDMS is transparent, for better visualization of the images shown in Figure 2, PDMS disks were color-coded by adding store bought food coloring dye into the mixture. A flat polyethylene sheet was placed on top, and the setup was cured at 70 °C for 2 h. After curing, the polyethylene sheet was removed and the PDMS disk was peeled from the plastic mold.

PAAm prepolymer with acrylamide to bis(acrylamide) (Bio-Rad) ratios of 7.5/0.24 wt % and 0.1% tetramethylethylenediamine (Sigma-Aldrich, T7024) was prepared in a centrifuge tube. PDMS disks were treated with 10% w/v benzophenone (Acros Organics, NJ) in diluted acetone (65% in water) for 60 s to swell and adsorb benzophenone into the silicone material. The samples were then rinsed with methanol and dried with nitrogen gas. 100 μL of freshly prepared 1% w/v ammonium persulfate initiator was added to 900 μL of PAAm prepolymer, mixed homogeneously using a vortex mixer, and added to the microwell molds. The PDMS disks were placed on top of the molds concentrically, and the setup was allowed to polymerize under UV light for 5 min to activate the surface-grafted benzophenone photoinitiator (Figure 2Bii). Upon gelation, the PAAm layer with patterned microwell features grafted onto the PDMS disk was carefully peeled from the mold to form the PAAm–PDMS composite disk. The PDMS base was thereafter perforated by punching holes around the polymerized PAAm using a standard 1 mm biopsy punch. The PAAm–PDMS disks were washed three times with PBS and incubated in PBS overnight on a shaker to leach out any excess unpolymerized toxins from the assembly. Prior to cell seeding, all the devices were kept hydrated by storing it under PBS at 4 °C. For hybrid integration, a ring magnet was aligned and placed under the device (as shown in Figure 2Biii), and collagen was added on top of the PAAm microwells to completely fill up the space within the PDMS gasket before attachment with the microfluidic channel assembly.

2.3 Cell Culture

Breast tumor cells MDA-MB-231 were used as the representative tumor cell line, and HUVECs were used as the representative endothelial model.46 HUVECs were cultured in endothelial growth media (ScienCell, 1001) supplemented with 5% fetal bovine serum (FBS), 1% penicillin–streptomycin (Gibco, USA), and 1% EC growth supplement (ScienCell, 1052). MDA-MB-231 cells were cultured in high glucose Dulbecco’s modified Eagle’s medium supplemented with 10% FBS and 1% Pen-Strep. Cells were cultured in standard cell culture flasks at 37 °C in a humidified incubator with 5% CO2 atmosphere. Immediately prior to the experiments, the cells were detached from the culture plates using Trypsin–EDTA (0.25%) and resuspended in complete growth medium to the desired concentration.

2.3.1 Microfluidic Endothelial Culture

Prior to EC seeding, all fabricated microchannels were exposed to germicidal UV light for 30 min and washed twice with PBS to remove any residual pluronic from bioprinting. Thereafter, the microfluidic channel was surface coated with poly l-lysine and stored overnight at 4 °C. The channel was washed twice with PBS to remove excess poly l-lysine. HUVECs were seeded onto the microchannel with a seeding density of 5 × 106 cells/mL. Excess HUVECs were removed after 2 h, then the integrated microfluidic channel Transwell insert was placed in a multiwell dish with culture media and incubated at 37 °C and 5% CO2.

2.3.2 Generation of Tumoroids

To generate tumor spheroids, the PAAm–PDMS disks were sterilized by exposure to UV light for 30 min, and then 100 μL of the tumor cell suspension (5 × 106 cells/mL) was gently pipetted onto the PAAm surface under sterile conditions. The setup was left undisturbed for 5 min to allow the cells to settle into the microwells by gravity. Thereafter, the PAAm microwells were gently rinsed with culture media to wash off the excess cells on the surface of the microwell plate. The resulting devices were stored in multiwell plates with culture media and incubated at 37 °C and 5% CO2 for 24 h to allow initial spheroid compaction.

2.3.3 Preparation of Collagen

To prepare the collagen matrix, type-I bovine collagen (Advanced Biomatrix, 5 mg/mL) was diluted to 1.5 mg/mL in PBS, 1× RPMI, and 10× MEM solution to obtain the final concentration. The acidic solution was neutralized with 1 M NaHC03 by titration based on the color of the indicator in RPMI. The prepolymerized collagen was kept in ice until further use. Immediately preceding the magnetic attachment of the microwell plate assembly devices with the endothelialized microfluidic channel, cold collagen gel solution was pipetted over the spheroids generated in the PAAm microwells to sufficiently fill up the space within the PDMS gaskets, and after attachment, the setup was allowed to incubate at 37 °C for 45 min for the collagen to gel.

2.4 Immunostaining

For assessing the intercellular adherens junctions, HUVECs were immunostained for vascular endothelial cadherin (VE-cadherin) using rabbit anti-VE cadherin monoclonal primary antibody (Invitrogen) and secondary goat antirabbit IgG (Abcam ab150116) antibody tagged with Texas red. Immunostaining for YAP/TAZ was performed using YAP1 polyclonal antibody (PA1–46189, Invitrogen) and Alexa Fluor 647 goat anti-rabbit IgG secondary antibody (A-21245, Thermofisher), and TAZ polyclonal primary antibody (PA5–58836, Invitrogen) was used in conjunction with Alexa Fluor 488 donkey anti-rabbit IgG secondary antibody (A-21206, Thermofisher). To quantify proliferation, we immunostained the HUVECs for Ki67, a common proliferation marker, using Ki67 recombinant rabbit monoclonal antibody (MA5–14520, Invitrogen) and Alexa Fluor 594 donkey anti-rabbit IgG secondary antibody (A-21207, Thermofisher). For immunofluorescent staining of the actin cytoskeleton, conjugated Alexa Fluor 488 phalloidin (A12379, Invitrogen) or Alexa Fluor 568 phalloidin (A12380, Invitrogen) was used depending on the assay.

HUVECs were initially washed in PBS and fixed in 4% (v/v) paraformaldehyde in PBS for 1 h at room temperature. The devices were then washed twice with PBS for 5 min each and permeabilized with 0.1% (v/v) Triton-X in PBS for 10 min at room temperature. After washing 2× with PBS for 5 min each, the ECs were blocked against nonspecific binding with 2.5% (v/v) goat serum in PBS for 1 h at room temperature. Cells were then incubated overnight at 4 °C with primary antibody in a 1:500 dilution of goat serum, washed and incubated with the 2.5% (v/v) goat serum for 30 min, and incubated with a 1:500 dilution of secondary antibody diluted in 2.5% goat serum for 1 h at room temperature. Samples were then thoroughly washed in PBS, and cell nuclei were counterstained using a 2 μg/mL DAPI (Invitrogen) in PBS for 30 min.

2.52.5 TEER Measurements

To quantify endothelial barrier integrity over longer term cultures, we monitored the permeability of the endothelium via daily TEER measurements and conducted end-point immunostaining for VE-cadherin at every 24 h of culture. For experiments that required TEER measurements, an open microfluidic channel configuration of the device was used for the endothelial culture. Electrical resistance across HUVECs was measured using the electrical cell sensor system, the Volt-Ω m (Millicell-ERS, Millipore, World Precision Instruments, Saratoga, FL, USA). TEER values were obtained for each day after the endothelial monolayer formation (days 1–6) and upon coculture with tumor cells (24, 48, and 72 h: days 7–9). The Volt-Ω m was calibrated before each use, and the electrodes were sterilized in 70% ethanol for 10 min and air-dried. The electrode containing the shorter arm was placed in the upper compartment within the channel, and the longer arm was placed in the lower well of a Transwell with enough media until constant measurement values were obtained. A Transwell polycarbonate membrane insert without cells was used as the blank, while the other wells containing cells served as treatment samples (n = 3). The TEER value (Ω·cm2) was calculated by the formula: [R sample (the average resistance of experimental devices) – R blank (the average resistance of blank devices)] × A (the area of the endothelial channel), and the resultant values were used to obtain the graphs.

2.6 Drug Testing

To screen interventional approaches for reversing the tumor-induced pro-angiogenic endothelial behavior, we first tested the efficacy of bevacizumab (Avastin), a humanized anti-vascular-endothelial-growth-factor-A antibody. Also, we targeted the Notch signaling pathway using nirogacestat, a selective γ-secretase inhibitor. Bevacizumab (HY-P9906, MedChemExpress, NJ, USA) was prepared to a concentration of 0.01 mg/mL in endothelial growth media. Nirogacestat (HY-15185, MedChemExpress, NJ, USA) used in the experiments was prepared to a concentration of 10 nM. The drugs were perfused into the endothelialized channel for the duration of the assay (24 h) using a syringe pump at a flow rate of 2 μL/h.

2.7 Imaging and Statistical Analysis

For both bright-field and fluorescent imaging, an inverted fluorescence microscope coupled with a CCD camera (Olympus IX-83) was used. Cell-Sens image analysis software (Olympus) was used for data acquisition. Image processing and analyses were performed using Image-J (NIH, USA). In all experiments, the formation of endothelial junctions under different experimental conditions was quantified by measuring the total length of the fluorescent marker exhibiting cell–cell junction lines. Unless otherwise specified, all the experimental data for EC parameters or different experimental conditions are expressed as mean ± standard error (SEM). Data sets were normally distributed, with similar variances between the compared groups. All statistical analysis was conducted using student’s t test or two-tailed one-way ANOVA analyzes with Tukey post hoc pairwise comparisons (Prism; GraphPad Software, La Jolla, CA). p-Values less than 0.05 were considered significant.

3 Results

3.1 Establishing the Tumor-Endothelial Coculture on M-TuVIC

3.1.1 Endothelialization of the Microfluidic Channel

Within 24 h of seeding and incubation, HUVECs were distributed uniformly and formed a confluent monolayer within the microfluidic channels. This was consistent for microfluidic channels with straight edges and also for serpentine channels that contained sharp corners and regions of high curvatures (Figure 3Ai). The HUVECs attached consistently to the porous membrane and spread to form confluent monolayers as determined by F-actin and DAPI staining (Figure 3Aii), consistent with previous reports.47,48

Figure 3 Enabling biomimetic coculture of the tumor and ECs using M-TuVIC A. (i) Confluent monolayer of DAPI-stained HUVECs inside a serpentine microfluidic channel showing uniform cell distribution along sharp corners and channel curvatures (scale bar 400 μm). (ii) Images of HUVECs stained for F-actin and nucleus demonstrate formation of monolayers on porous microfluidic channels after 24 h (scale bar 75 μm). (iii) Increase in the TEER measurements conducted across HUVECs cultured on an open microfluidic channel membrane over 10 days. Peak TEER was observed on day 7 of the culture without any significant decrease in the TEER values thereafter (data presented as mean ± SEM for n = 3 samples). (iv) Images of HUVECs at days 1 and 7 of culture immunostained for VE-cadherin (red) and counterstained with Hoechst (blue; scale bar 100 μm). B. (i) Images of hybrid integration of ECs and arrayed tumor spheroids. For visualization, HUVECs were stained with DAPI (blue; scale bar −600 μm). (ii) Fabrication of different microfluidic channel designs perfused with fluorescent dextran (red) on the porous membrane, aligned, and integrated with tumor spheroids (green; scale bar 200 μm). .

The open microfluidic channel device structure facilitated the use of standard chopstick probes for the TEER measurement. We found that TEER values increased with the culture duration and reached a maximum (∼105 Ω·cm2) on day 7 (Figure 3Aiii). The results from immunostaining also showed an increased localization of VE-cadherin at the cell–cell junctions after 7 days of culture compared to a 24 h culture (Figure 3Aiv), corroborating that the increase in TEER corresponds to the elaboration of EC–cell junctions. No significant difference was observed in the adherent junction length per unit area after 72 h of peak TEER value, indicating the sustenance of functionally tight cell–cell junctions after 7 days of the HUVEC culture within the channel.

3.1.2 Hybrid Integration of the Tumoroids with the Endothelial Monolayer

The grafting of PAAm onto the perforated PDMS disks during polymerization was consistent and robust. PDMS–PAAm composite microwell disk assembly was easy to handle during the tumoroids culture, and we were able to consistently generate an array of 3D tumor spheroids using the method described in Section 2.2.2. For tumor–vascular interface studies that do not require integration of the endothelialized microfluidic channel with the tumoroids, microwell plate assembly required alignment, which was achieved by placing a ring magnet under the PDMS disk over a flat surface and positioning the Transwell assembly (containing the microfluidic channel and a ring magnet placed inside) from the top, close to the PDMS disk so as to enable instantaneous magnetic attachment. To demonstrate this, we created tumor–vascular interfaces by aligning and integrating straight and serpentine microfluidic channels seeded with ECs with tumoroid arrays (Figure 3Bi). Other geometric features of the vasculature such as diameter, curvature, and branching were recapitulated by fabricating different microfluidic channel configurations similar to the framework of the vasculature-like networks49 and integrating the channels with fluorescently labeled tumor spheroids, with controlled respective positioning along the imaging plane (Figure 3Bii). Regardless, this setup could always accommodate for engineering tumor–vascular interface coculture models that do not require the confinement of the ECs inside microfluidic channels but could still benefit from a higher analysis throughput, enabled by the array of spheroids magnetically attached to an endothelialized porous membrane (Supporting Information Figure S3). We were able to image HUVECs even when tumoroids were in the light path by detaching the tumor microwell plate from the assembly and attaching them back if required (Supporting Information Figure S3). These results demonstrate that the M-TuVIC enables the biofabrication of a tumor–vascular interface with wide range of vascular geometrical configurations, and overall, the magnetically enabled hybrid-integration approach ensured adequate spatial compartmentalization of the cultures within the integrated assembly while providing maneuverability to precisely align the different subcomponents of the M-TuVIC as a stacked 3D coculture model.

3.2 Study of Tumor-Mediated Endothelial Dysregulation

During tumor angiogenesis, tumor cells induce structural and functional changes in nearby blood vessels through the activation of previously quiescent ECs into a dysregulated phenotype.13 Previous works have demonstrated that tumor cells modulate EC spreading dynamics and proliferation rate,50,51 and so to determine whether the tumor cells were capable of signaling to the HUVECs in the M-TuVIC device, we established a process flow for the coculture assays (Supporting Information Figure S4) based on the device characterization presented in Section 3.1. HUVEC proliferation at different regions of the microchannel relative to the location of a tumoroid was observed using an M-TuVIC configuration, wherein the spheroids were arranged linearly spaced apart by ∼1.4 mm, and the width of the endothelial channel was greater than the diameter of the tumoroids (schematically shown in Supporting Information Figure S5). ∼60% of HUVECs in the closest proximity (less than the spheroid diameter, d, i.e., ∼350 μm) to the tumoroids showed positive Ki67 signal (Figure 4Ai). The number of Ki67+ decreased with an increasing distance from tumoroids (Figure 4Ai). A significantly greater subpopulation of the HUVECs located within d showed a greater expression of Ki67 (p < 0.05) than the cells present within the next closest range of d ∼ 2d. However, no significant difference was observed in the count of Ki67+ HUVECs when situated at any distance greater than 2d (i.e., ∼700 μm) from the tumoroids (Supporting Information Figure S6), suggesting that any tumor-endothelial coculture must take into account the proximity of the cultures for measuring the endothelial sensitivity response to the presence of tumor. Furthermore, regions with a greater proportion of Ki67+ HUVECs were significantly (p < 0.001) more densely packed (Figure 4Aii) as compared to a control endothelial monoculture.

Figure 4 Proliferation of the HUVECs located at different distances from the tumor (expressed in terms of the spheroid diameter d) was assessed by the ratio of the average number of Ki67-positive cells to total cells in 9 random high-magnification fields of view. A significant difference in Ki67-positive HUVECs was observed in the vicinity of tumor vs away from the tumor. The data were expressed as the mean ± SEM of three independent experiments (N = 3; *p < 0.05). (ii) Cell density per field of view of the HUVECs in proximity to the tumor was assessed by counting the total number of DAPI-stained nuclei, which showed a significant increase within 24 h of coculture with the tumor spheroids. (B) (i) Immunofluorescent characterization of HUVECs after 24 h of coculture with tumor spheroids demonstrates loss of the well-developed network of adherens junctions (VE-cadherin, red) and increased cell density (Hoechst nuclear stain, blue) compared to the control monoculture (Scalebar represents 250 μm). This was quantified using (ii) endothelial adherens junction length measured per unit area (VE-cadherin stained), which is significantly reduced with tumor coculture. (iii) Significant decrease in nuclear spread area, which demonstrates significant changes in cell morphology as a function of tumor coculture (iv) TEER was measured across the ECs cocultured with the tumor and showed a significant decrease over the duration of the coculture compared to endothelial monoculture over the same time period. Data is presented as mean ± SEM for a minimum of N = 3 experiments and a minimum of n = 50 for single-cell measurements over at least three fields of view presented in panel B(i) with *p < 0.05, **p < 0.01, and ***p < 0.001, using two-tailed one-way ANOVA with Tukey post hoc comparisons.

End-point immunofluorescence imaging of endothelial morphology and markers of adherens junction formation between the ECs showed that the presence of tumoroids disrupted the endothelial lining. We observed a dramatic change in VE-cadherin network formation (Figure 4Bi), with statistically significant disruptions to the VE-cadherin network with increasing duration of the coculture (Figure 4Bii), accompanied by the changes in cell shape and density indicated by a significant reduction (p < 0.001) in the nuclear spread area (Figure 4Biii) even within the first 24 h of the hybrid integrated coculture. The loss of VE-cadherin corresponded with a significant (p < 0.05) drop in the TEER values (Figure 4Biv), in comparison with the respective monoculture control at the same time point. Continuity in the trend of these tumor-endothelial characteristics with prolonged coculture, as observed at the 48 and 72 h time points (Supporting Information Figure S7), further validated our 24 h observations, and after 72 h, almost negligible levels of VE-cadherin were detected at cell–cell junctions. The TEER values corroborate the loss of endothelial permeability,24,52 and these results are consistent with other studies demonstrating increased endothelial proliferation,53 adherent junction loss, and drop in TEER in the presence of tumors, re-emphasizing the well-established importance of cell and nuclear morphology on critical cell functions including apoptosis,54 endothelial–mesenchymal transition,55 and other fundamental processes that occur during developmental, homeostatic, and pathological angiogenesis. Thus, our hybrid integrated coculture establishes the biomimetic capability of the M-TuVIC to model these characteristic tumor-vasculature behaviors and recapitulate the necessary details.

3.3 Pro-angiogenic Endothelial YAP/TAZ Nuclear Localization

YAP and TAZ, the main effectors of the Hippo pathway, have emerged as important players in cancer biology and therapy response,56 and the intracellular localization of YAP/TAZ is a key determinant in the regulation of their activity and their roles in signal transduction. VE-cadherin, present along the adherent junctions of the ECs, is connected to the actin cytoskeleton through phosphorylated YAP/TAZ.57 It has been reported that activated endothelial YAP/TAZ in turn induces a downstream cyclic transcriptional program which further regulates proliferation, actin cytoskeleton contractility, and cell adhesion, leading to endothelial dysregulation and collective cell migration.58 Therefore, we hypothesized that tumor-mediated loss of VE-cadherin along the endothelial junctions would also result in the redistribution of the YAP/TAZ, and so to test this hypothesis, we immunostained ECs for YAP and TAZ.

In monoculture conditions, YAP/TAZ was colocalized with F-actin-stained regions of the cell, while in coculture, YAP/TAZ was colocalized with DAPI (Figure 5i). To further quantify these differences, we plotted the normalized relative line intensity of the YAP/TAZ expressions across a single EC structure between cell-to-cell junctions identified visually by using the fluorescent images (Figure 5ii). In the absence of tumoroids, sharp peaks in the endothelial YAP/TAZ were observed outside the nuclei, while coculture with tumoroids results in greater YAP/TAZ intensity within the nuclei, as well as several mini peaks that are not associated with the cytoplasm (Figure 5ii). We further observed ∼99% overlap in the fluorescence signal between YAP and TAZ (Supporting Information Figure S8), showing that YAP and TAZ colocalize under these conditions, as expected.59 Overall, we observed a statistically significant reduction (p < 0.001) in cytoplasmic YAP/TAZ distribution and a simultaneous statistically significant increase (p < 0.01) in the YAP/TAZ nuclear translocation under tumor-endothelial coculture (Figure 5iii), consistent with previous observations that showed elevated nuclear localization of YAP and TAZ in the ECs under malignant conditions.58,60,61

Figure 5 YAP/TAZ expression in ECs (i) Immunostaining the HUVECs for YAP (purple), TAZ (green), nuclei (DAPI-blue), and F-actin (red) shows the YAP/TAZ nuclear translocation under tumor coculture as compared to cytoplasmic YAP/TAZ expression under control monoculture condition. Scale bar represents 100 μm. (ii) Line intensity of normalized fluorescence expression is plotted from the sectional lines (shown as red dashed) across the fluorescence images of a single EC for control and coculture conditions (scale bar represents 50 μm) (iii) Data collected from the immunostaining shows a significant decrease in the mean cytoplasmic fluorescence intensity and a simultaneous increase in the mean nuclear fluorescence intensity of the endothelial YAP/TAZ under tumor coculture (data presented as mean ± SEM for a minimum of N = 3 experiments and a minimum of n = 50 for single-cell measurements over at least three fields of view presented in panel A(i) with **p < 0.01 and ****p < 0.0001, using two-tailed one-way ANOVA with Tukey post hoc comparisons).

3.3.1 Demonstration of Antiangiogenic Drug Testing

Endothelial YAP/TAZ activation is important for the formation of new blood and lymphatic vessels during developmental angiogenesis.61 However, research into the mechanisms that regulate YAP/TAZ activity in tumor ECs, and the upstream signals from the TVME that control endothelial YAP/TAZ activation in pathological angiogenesis are still evolving.58,60,62 Within a TVME, the tumor cells are a rich source of vascular endothelial growth factors (VEGFs), which function as a potent stimulant of the pro-angiogenic endothelial switch that regulates all the key steps of the angiogenic process, including EC proliferation and migration.63 Therefore, we first tested the efficacy of bevacizumab (Avastin), a U.S. Food and Drug Administration (FDA)-approved anti-VEGF drug, on endothelial normalization. Thereafter, we evaluated the efficacy of nirogacestat, a reversible, noncompetitive gamma secretase inhibitor (GSI) that selectively blocks Notch signaling.64

After hybrid integration, bevacizumab was continuously infused into the microfluidic channel for the duration of the coculture (24 h). Within the chosen concentration range, bevacizumab treatment resulted in a significant sequestration of the YAP/TAZ from the nuclei into the endothelial cytoplasm (Figure 6Ai), and the endothelial adherens junction integrity was sustained even under 24 h coculture (Figure 6Aii), as quantified with a significantly greater VE-cadherin expression at the cell–cell junctions (Figure 6Aiii) upon drug treatment. These data are demonstrative of the capacity of anti-VEGF-A blocker in regulating the pro-angiogenic tumor-endothelial behavior and further suggest that the changes in HUVECs associated with coculture are driven in part through VEGF-mediated signaling.

Figure 6 Drug-mediated reversal of tumor-endothelial behavior. (A) Bevacizumab: (i) immunostaining of bevacizumab-treated ECs under coculture shows distribution of YAP (purple)/TAZ (green) in the cytoplasm away from the nuclei (DAPI-blue); (ii) immunofluorescent characterization of the ECs upon coculture with tumor demonstrates the sustenance of adherens junction network (VE-cadherin, red) with less densely packed cells (Hoechst nuclear stain, blue) after 24 h of treatment with bevacizumab, compared to the control (no-drug treatment) condition. This was quantified by (iii) adherens junction length measured per unit area showing significantly higher VE-cadherin network with bevacizumab treatment (data is presented as mean ± SEM for a minimum of N = 3 experiments and a minimum of n = 50 for single-cell measurements over at least three fields of view presented in the representative images, ***p < 0.001) (scale bar = 50 μm). (B) Nirogacestat: quantification of YAP and TAZ localization within HUVECs with and without (control condition) treatment with nirogacestat, with N < C denoting less YAP and TAZ in nucleus than in cytoplasm; N = C, similar levels of YAP and TAZ in cytoplasm and nucleus; N > C, more YAP and TAZ in nucleus than in cytoplasm (data is presented as mean ± SEM for a minimum of N = 3 experiments and a minimum of n = 50 for single-cell measurements over at least three fields of view presented in Supporting Information Figure S9).

Using a similar drug testing approach, 10 nM nirogacestat was infused into the endothelial channel over 24 h of tumor-endothelial coculture. For the quantification of YAP and TAZ localization, we measured the fluorescence intensity of the YAP/TAZ within the nuclei and cytoplasm of the ECs.65 Nirogacestat is the first Notch GSI to begin early phase clinical trials with no concerning safety signals66,67 and was recently approved by the FDA for treatment of desmoid tumors and is in further clinical development as an antiangiogenic agent for advanced triple negative breast cancer. Qualitatively, for the ECs treated with either bevacizumab or nirogacestat, YAP/TAZ appeared to be more uniformly distributed within the cytoplasm (Supporting Information Figure S9) compared to clustered cytoplasmic localization of the endothelial YAP/TAZ under tumor coculture without drug treatment (Figure 5i). This behavior could be indicative of the YAP/TAZ activity within the ECs60 as a result of nondirectional sequestration and redistribution of YAP/TAZ from the nuclei into the cytoplasm of activated ECs during the process of YAP/TAZ nuclear translocation in the presence of tumors. We further observed that nirogacestat treatment induced a significantly decreased colocalization (p < 0.001) of YAP/TAZ with endothelial nuclei compared to untreated conditions (Figure 6B), suggesting that modulation of the Notch signaling pathway using nirogacestat could function as an antiangiogenic strategy by regulating the YAP/TAZ-mediated cytoskeletal dynamics of the tumor-activated ECs.

4 Discussion

In this work, using our M-TuVIC, we have successfully demonstrated the early stage tumor–vascular interactional dynamics16 defined by tumor-stimulated proliferative and permeable endothelium, pro-angiogenic organization through adherens junction loss, endothelial proliferation, and YAP/TAZ nuclear localization in a time-dependent manner. Based on this endothelial signaling, we have further demonstrated the combined utilities of the M-TuVIC platform for early stage drug screening through different signaling pathways of vascular normalization strategies.

Tumors are highly related to the blood vessel-dominated microphysiological environment,68 and therefore, integrating vasculature into the existing 3D tumor models offers several advantages that would open up the feasibility of conducting a plethora of in vitro studies specifically recapitulating critical stages of the pathophysiological events that occur at the tumor–vascular interface. However, such in vitro experiments could often be encountered by problems typically faced within a multicell culture environment, which challenge assay protocols as well as the phenotype and output for analyses. For instance, within typical mixed coculture models or enclosed microfluidic cell culture systems, isolating the physical contact of multiple cells or tissue types during multistage cell seeding is always cumbersome, and our results show that mere proximity of the tumor cells to the ECs even without direct physical contacts could be detrimental to the sustenance of a functional endothelial barrier marked by the formation of mature adherens junctions. Consequently, basic models of the tumor vasculature created by a simplified mixed coculture of tumor and ECs would neither be representative of their in vivo-like functionalities nor be able to provide the individual cell populations with biochemical signals in a physiologically relevant context within the coculture. Using the M-TuVIC, we show the feasibility of segregated coculture, wherein the functionalities of the individual subpopulation of cultures can be continuously monitored up until maturation before introducing them into a biomimetic coculture format whenever required.

Whereas the advantages of on-chip vascularization with integrated microfluidics69 is well-known, the improvisation offered by our rapid-prototyping friendly approach enabled by magnetic integration facilitates the engineering of cellular microenvironments with high degrees of spatiotemporal precision, and on-the-fly customizability of the platform to accommodate application-dependent design changes to the different subcomponents. Any required layout of the tumoroids’ arrangement could be easily achieved by modifying the design of the 3D printed mold to transfer the pattern onto the hydrogel during the fabrication of the microwells. Likewise, a broad range of vasculature contours can also be obtained by spontaneous changes to the microfluidic channel design during the initial stages of bioprinting. The magnetically enabled hybrid integration approach of the M-TuVIC device also offers the advantage of isolating and retrieving the individual cultures at any time point, thereby enabling closer post-assay analyses through nondestructive detachment of the ensembles for routine procedures such as TEER measurements or immunostaining. The importance of YAP/TAZ function for vascular development has been studied by several groups using mouse models,57,62 but surprisingly, research works using organ-on-chip technologies to demonstrate the pro-angiogenic modalities of YAP/TAZ intercellular redistribution are scant, possibly due to the limitations with such technologies, as described earlier. Herein, we show that YAP/TAZ nuclear localization can be used as a critical parameter to study tumor-endothelial behavior in vitro, and such high-resolution examination of the ECs for intercellular YAP/TAZ redistribution reiterates the significance of the mechanisms by which endothelial YAP/TAZ signaling could achieve combinatorial control of gene expression and cooperation of other signaling pathways during developmental angiogenesis.56 In general, magnetic hybrid integration provided robustness to maintain the integrated assembly without mechanical disturbances associated with regular procedures, such as handling the device during transfer of a cultured vessel or repeated movement of the microscope stage for imaging. Thus, our unique hybrid-integration approach, adopted to engineer a biomimetic tumor–vascular interface, facilitated the implementation of a multipart coculture assay to reveal certain critical biological phenomena while limiting the overall size and complexity of operation. By applying simple design modifications to the current format, the M-TuVIC could be immediately applied to further study the influence of other physical and biological microenvironmental parameters at the tumor–vascular interface, such as fluid shear stresses, vascular curvatures, tumor size, endothelial insult mechanisms, etc.

One of the key challenges, though, for advancing the capabilities of the M-TuVIC device platform is assay-specific optimization and validation of all the associated design, development, and testing protocols. Complex microphysiological events that occur at the tumor–vascular interface, such as angiogenesis and tumor extravasation, could be a resultant of a cascade of several different phenomena involving multiple cellular and tissue-level components within a TVME. To be replicated within an in vitro setting, this may necessitate unique media requirements and combinations of growth factors, introduction of other components of the TVME68 into the existing setup, together with a substantially longer coculture adequately controlling for the complementary responses or changes in the behavior of each of the individual cell types and their signaling cross-talks,70 which would call for extensive and careful characterization of all the entities involved. Furthermore, increasing the complexity of the assays could also require the incorporation of additional readouts for data quantification, in which case the current data acquisition approach through immunofluorescence imaging using planar microscopes may hit a limitation in terms of resolution, optical saturation, and imaging depth for an arrangement like the M-TuVIC that consists of 3D stacked heterogeneous cocultures built in with optically interfering materials. This would then require a need to move toward more rigorous 3D imaging and reconstruction techniques, adding more complexity to the overall design of experiments. Nevertheless, the design adaptability of the M-TuVIC provides confidence that with appropriate controls and optimization procedures, the magnetically hybrid integrated multilayer coculture strategy could provide a potentially scalable approach to effectively include advanced genomic or transcriptomic level analysis of the heterogeneous cocultures that could help with the quantification of complex, physiologically relevant assays and development of suitable therapeutic strategies. Thus, the M-TuVIC is not only useful for studies at the tumor–vascular interface but also, in general, could be extrapolated for the development of experimental platforms requiring 3D cocultures of vascular disease models and segregated cell–cell interfaces, thereby providing a versatile and defined tool for benchtop investigations of early stage drug discovery processes.

5 Conclusions

This work reinforces the importance of incorporating vasculature into existing models of the tumor microenvironment as a critical factor for gaining unique insights into significant biological events related to tumor development and early stage pro-angiogenic vascular dysfunctions. Our M-TuVIC model provides a viable solution to mimic the physiological TVME, and our results demonstrate the utility of the device platform for probing further into the organization of tumor vasculature and for conducting antitumor drug testing by targeting tumor-induced pro-angiogenic endothelial dysregulations, as well as the feasibility of drug-mediated reversal by modulation of signaling pathways previously unreported for in vitro antiangiogenic drug screening. The overall fabrication procedure of the device is rapid and simple and does not require any specific engineering skill set, making this device platform accessible to a broad spectrum of users. We believe that the M-TuVIC platform can provide better clinical predictions by bridging the gap between proof-of-concept studies and a broader implementation in potentially high-throughput, scalable biomimetic drug screening approaches for different disease conditions.

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acsami.4c01766.Schematic of fabrication of the composite polymer-hydrogel microwell disk assembly, schematic of sacrificial bioprinting of the microfluidic channel on porous membrane, representative image of tumor-endothelial coculture, tumor-endothelial coculture assay process flow, design layout for endothelial proliferation assays, fluorescent images of EC proliferation, qualitative and quantitative representation of tumor-endothelial behavior, endothelial YAP/TAZ distribution, and YAP/TAZ distribution in nirogacestat-treated ECs (PDF)

Supplementary Material

am4c01766_si_001.pdf

The authors declare no competing financial interest.

Acknowledgments

The authors thank Dr. Roni F Rayes from McGill University, Montreal, Canada, and Dr. Fei Xing from Wake Forest School of Medicine, USA, for useful discussions on this work. The work was supported by the National Institute of Health (5SC2GM136523 to A.C.) and the American Heart Association (CDA857738 to W.J.P.). The authors would like to thank the financial support of the NC Translational and Clinical Studies (NC TraCS 550 KR282124 to S.S.), US Department of Defense (W911NF2120265 to R.A.), and National Institutes of Health (T32HL69768 to C.G.). The work was conducted per the guidelines of the Institutional Biosafety Committee (IBC#23-01), NC A&T State University.
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