Lab on a Chip

We demonstrate a new sample-loading strategy ('spot-loading') for Spores-on-a-Chip microfluidic sensors that enables multiplexed experimentation. The previous limitation of one biological sample per device is overcome through controlled sample loading at an intermediate step of the chip fabrication process. As a proof-of-concept, we use dual spore chips to compare the germination behaviours of two spore strains in two distinct microenvironments.

Tumor models that recapitulate 3D architecture are essential for understanding how cellular organization and microenvironmental interactions govern therapeutic response in human cancers. Here, we developed a microfluidic microphysiological system that enables controlled and scalable culture and drug testing of non-small cell lung cancer spheroids and patient-derived organoids. The platform integrated U-shaped microwells with dual-channel loading to support de novo spheroid formation, efficient trapping of pre-formed spheroids, and loading of intact organoids with reduced size heterogeneity. Tumor spheroids and organoids maintained high viability and structural integrity during long-term on-chip culture, and constrained microscale confinement produced ellipsoidal geometries that deviate from idealized spherical assumptions. Baseline genotype-dependent responses to KRAS G12C and EGFR inhibitors were preserved across agarose and microfluidic formats, establishing a validated reference state. Building on this baseline, fibroblast- and endothelial-derived cues consistently attenuated responses to targeted therapies across conditioned media, mixed co-culture, and spatially organized configurations. Resistance phenotypes converged on a dominant role for paracrine signaling, while increasing architectural complexity primarily enhanced morphological fidelity rather than altering therapeutic response. These findings establish a microphysiological framework that decouples tumor-intrinsic drug sensitivity from microenvironment-mediated modulation, enabling the systematic evaluation of paracrine resistance mechanisms in NSCLC.

Metastatic dissemination is initiated by tumor cells interpreting spatially organized biochemical and biophysical cues that remain difficult to reproduce using conventional migration assays. Here, we developed a computationally guided metastasis-on-a-chip (MET-on-a-chip) platform based on the concept of the Minimally Functional Unit (MFU), in which only the biological components required to answer a defined experimental question are incorporated. The platform consists of two independent culture chambers connected through an array of confined microchannels that permits diffusion of soluble factors while constraining tumor cell migration. Rather than relying on empirical optimization, finite-element COMSOL simulations were first used to predict molecular transport, define growth factor loading conditions, identify biologically relevant exposure regions, and guide the rational design of the microfluidic assay. Computational predictions were experimentally validated using 70-kDa FITC-dextran diffusion and VEGF release studies, confirming the formation of stable spatial concentration gradients across the microfluidic platform. The simulations further demonstrated that both growth factor loading and cell positioning relative to the predicted gradients critically influenced assay performance, leading to the optimization of the platform through spatial reconfiguration of the tumor compartment. Using the optimized configuration, we compared the migratory responses of neuroblastoma, Ewing sarcoma, and osteosarcoma cells to vascular (VEGF-A165) and lymphatic (VEGF-C) chemotactic cues. VEGF-C significantly increased migration through the microchannel array in Ewing sarcoma and osteosarcoma cells, whereas VEGF-A165 produced no significant effect. In contrast, neuroblastoma cells exhibited minimal migration under either condition, revealing tumor-specific differences in responsiveness to VEGF signaling. Together, these findings establish a computationally guided workflow for the rational design of metastasis-on-a-chip assays, in which predictive modeling informs experimental design before biological validation. By substantially reducing empirical trial-and-error while enabling quantitative control over growth factor exposure, this strategy provides a robust framework for developing minimally functional microphysiological systems capable of dissecting individual steps of the metastatic cascade under experimentally defined conditions. Translational Impact StatementMetastatic dissemination remains one of the greatest clinical challenges in pediatric oncology, yet experimental models capable of quantitatively evaluating early migratory events remain limited. The computationally guided MET-on-a-Chip workflow presented here provides a human-relevant platform in which soluble microenvironmental cues can be systematically investigated under controlled and predictive conditions. Although demonstrated here using VEGF-A165 and VEGF-C, the platform can be readily adapted to study virtually any chemotactic factor, cytokine, extracellular vesicle population, or therapeutic candidate involved in metastatic dissemination. The modular MFU design allows biological complexity to be incorporated progressively as dictated by the scientific question, providing a flexible framework for future applications. In the longer term, this workflow could be combined with patient-derived tumor cells, organoids, or biopsy material to investigate patient-specific metastatic behavior and evaluate anti-metastatic therapeutic strategies in a personalized setting. Beyond identifying pro-migratory signaling pathways, the platform may serve as a preclinical tool to prioritize compounds capable of preventing tumor cell dissemination before evaluation in more complex animal models or clinical studies.

Inflammation-driven increases in nociception are prominent in pain pathologies associated with intervertebral disc (IVD) degeneration yet are difficult to model in vitro. Since neurons are exposed to multi-modal stimuli in vivo, it is critical for these exposures to be conserved in an in vitro test system. We developed a polydimethylsiloxane (PDMS)-based microfluidic platform to interrogate peripheral sensory neurons (SNs) in the presence of conditioned media from nucleus pulposus cells from the degenerated IVD, to model a potential impact of IVD cells secretome on pain sensing. Our platform enables controlled perfusion of cell-derived biochemical cues alongside a defined homogeneous electric field (EF) and supports real-time optical analysis. Computational modeling, fluid perfusion experiments, and conductivity measurements confirmed stable fluid transport and tunable homogeneous EF generation within the device. As proof of concept for neuronal stimulation, neuroblastoma (N2a) cells loaded with a fluorescent Ca2+ indicator exhibited a 56% increase in Ca2+ transient activity when exposed to media from degenerated IVDs, concomitant with increased IVD-derived IL-1{beta} production. Importantly, EF-stimulated Ca2+ transients increased in SNs derived from human induced pluripotent stem cells when exposed to conditioned media from primary human IVD cells, demonstrating the translation of this model system to human cells. Together, these results establish a versatile platform that enables controlled and simultaneous exposure to biochemical and electrical stimuli to quantify inflammation-driven peripheral neuronal hyperexcitability in tissue-neuron crosstalk.

Immune-epithelial interactions govern the initiation and progression of airway diseases, yet their heterogeneity is difficult to capture using existing in vitro models. Although conventional Transwell and lung-on-chip systems reproduce airway compartmentalisation and permit epithelial-immune interactions, they lack the spatial and analytical resolution needed to visualise dynamic immune behaviour during infection. Here, we present the "Single Cell resolved Airway-Immune Recruitment" (scAIR) platform designed to interrogate immune-epithelial interactions during airway infection. The scAIR device features a modular central chamber accommodating a Transwell insert with primary airway epithelial cells (AECs) pre-differentiated under air-liquid interface (ALI), flanked by immune compartments connected through a precision-patterned microchannel array. This architecture enables real-time single-cell imaging of immune cell migration while preserving epithelial physiology. The scAIR device coupled with a machine learning analysis (MLA) pipeline enables automated tracking and quantification of individual immune cell speed, direction, and behavioural heterogeneity. Using this platform, respiratory syncytial virus (RSV) infection is modelled to generate a type 1 inflammatory airway epithelium that drives neutrophil recruitment. TNF-alpha neutralisation with adalimumab reveals distinct migratory behaviours that are obscured by population-averaged measurements. This integrated platform quantifies airway immune responses during infection and therapeutic modulation, enabling mechanistic studies, drug evaluation, and precision modelling of airway inflammation.

Organoids-on-chip combine the 3D complex microenvironment and cellular composition of organoids with microfluidic flow, increasing nutrient-waste exchange and mimicking the contributions of in vivo interstitial and vascular flow. However, the widespread adoption of organoid-on-chip platforms is limited by the lack of incubator-friendly flow control systems. Existing approaches often rely on commercially available syringe or peristaltic pumps, but these are bulky, lack scalability, and present a significant barrier for clinical translation. To circumvent these issues, we present the Compact Active Perfusion Standalone Organoid-on-Chip (CAPS-OC) platform, a fully integrated and battery-powered microfluidic system capable of culturing organoids in active media flow. To achieve this, we introduce a novel low-power mechanism of pressure pulse generation using an off-the-shelf compact rotary actuator (CRA), and package it into a compact electromechanical assembly. This assembly provides timed pneumatic inputs to achieve programmable control of membrane-based peristaltic pumps, with [~]100 {micro}L/hr dynamic range achieved on a custom microfluidic organoid chip. We biologically validated this system by culturing pancreatic cancer organoids derived from a KrasLSL-G12D/+; Trp53LSL-R172H/WT; Pdx1-Cre (KPC) genetically engineered mouse model. We found that our chip enhances proliferation and helps sustain a population of larger (>150 {micro}m) organoids compared to standard dome based static culture. Additionally, through immunostaining, we observe that KPC organoids cultured in the chip show more aggressive PDAC phenotype with reduced nuclear expression of GATA6, whereas organoids in static culture retain less aggressive classical- like subtype. Finally, testing of a RAS inhibitor drug, daraxonrasib, on the KPC organoids on chip showed size-based sensitivity elucidating the impact of active perfusion on the drug diffusion kinetics. Altogether, we establish CAPS-OC as a valuable tool for the bioengineering community and for clinically translating organoid model systems.

Circulating tumor cells (CTCs) are valuable biomarkers for cancer diagnosis and monitoring, yet their isolation from blood remains challenging due to their phenotypic heterogeneity and rarity. Label-free microfluidic technologies offer a promising alternative to affinity-based approaches by exploiting intrinsic biophysical differences between cell types. Here, we developed a microfluidic platform for label-free cell separation based on insulator-based dielectrophoresis (iDEP). The microfluidic device employs an array of triangular insulating structures that generate strong electric field gradients in response to an externally applied alternating current (AC) electric field, enabling selective isolation of breast cancer cells from blood cells based on their dielectric properties. Hydrodynamic focusing is used to confine the sample stream and precisely control cell trajectories within the separation region. Numerical simulations were performed to optimize the electric field distribution and fluid flow characteristics within the device. Experimental validation using breast cancer cell lines (mesenchymal-like MDA-MB-231 cells and epithelial-like MCF-7 cells) spiked into peripheral blood mononuclear cells (PBMCs) demonstrates selective dielectrophoretic deflection of cancer cells while PBMCs largely follow the central streamline. The platform achieves recovery rates exceeding 98% and a separation purity above 65% within the optimized operating conditions. The proposed system provides a simple label-free approach to separate heterogeneous cell populations and represents a promising tool for microfluidic liquid biopsy enrichment applications.

Control of fluids is a hallmark of microfluidic systems and fundamental for the successful application of microfluidic devices. Trigger valves use geometric features to autonomously control the release of fluids in microfluidic devices. Our previous work has adapted geometries used in closed trigger valve systems to enable use in open systems, allowing for open microfluidic devices with up to three trigger valves. Here, we focus on the parallel co-flows produced by sequential release of trigger valves and present a model that predicts their layer widths as a function of the geometric characteristics of the different side channels of each trigger valve. We show layered co-flows with widths as low as 50 microns. Additionally, we expand the use of trigger valves in open microfluidic devices by incorporating 1) varied step heights, 2) devices with up to seven trigger valves, and 3) use of varied fluids and plastics. To validate the implementation and use of these trigger valves in open systems, we have developed a theoretical framework to compare predicted outcomes (i.e., fluid travel distance, velocity, and layering width) with our experimental values. This theoretical work offers applications in various fields, including hydrogel patterning for 3D cell culture, organ-on-a-chip models, at-home sample preparation, and autonomous microfluidic systems for biosensing.

Tracking suspended cells over multiple time points at the single-cell level remains challenging because existing flow-based methods cannot preserve cell identity while maintaining high throughput. Here, we present RASPBerry, a hydrogel-based spatial barcoding platform for time-lapse flow cytometry. RASPBerry generates unique barcodes by randomly co-encapsulating fluorescent beads with individual cells in hydrogel droplets, eliminating the need for predefined barcode patterns or specialized optical instrumentation. We integrate RASPBerry with acoustofluidic imaging flow cytometry to enable time-lapse imaging flow cytometry of suspended cells. The platform identifies more than 17,000 hydrogel droplets with 99.8% matching accuracy. We further demonstrate time-lapse tracking of more than 10,000 suspended cells and quantify stress-induced nuclear morphological changes in more than 5,000 individual cells. RASPBerry provides a simple, scalable, and broadly accessible strategy for time-lapse imaging flow cytometry, expanding the capability for dynamic single-cell analysis of suspended cells.

Sensitive and low-cost protein biomarker detection is critical for disease diagnosis. Advanced microfluidic systems can generate miniature reaction compartments for a high-sensitivity assay. However, these platforms often require external instruments, skilled operators, and complex setups. Here, we develop a Lab on a Capillary (LabCap) platform that integrates photopatterned hydrogel rings within a glass capillary using a reconfigurable stop-flow lithography system. During sample loading and unloading steps, nanoliter-scale aqueous droplets (torodrops) are spontaneously formed around the hydrogel rings, creating isolated reaction compartments without the need for external instruments or an immiscible oil phase. The LabCap platform enables quantitative detection of clinically relevant biomarkers, including C-reactive protein (CRP) and N-terminal pro-B-type natriuretic peptide (NT-proBNP). By adjusting the incubation protocol, assay speed and sensitivity can be tuned to meet different analytical requirements. A periodic medium exchange protocol enables biomarker detection at concentrations as low as 1 ng/mL, whereas prolonged static incubation extends detection to 0.1 ng/mL. In addition, LabCap offers practical advantages, including low fabrication cost (< EUR 1 per device), low reagent consumption (<100 microlitres per assay step), and minimal wash-buffer usage (1 mL). These results demonstrate that LabCap is a simple, cost-effective, and versatile platform for biomarker detection.

The role of spatial structure in microbial ecology and evolution is increasingly recognised and investigated, often with agar plates as a template for a spatially structured environment. While convenient, agar plates do not allow for the spatial and temporal control microbiologists have become accustomed to in the field of microfluidics with its tight environmental control for single cells and small populations, holding back research on surfaces and at larger length scales and population sizes. To close this gap, we developed a novel device with an agar sheet sealing indented channels through which media perfuses. As proof of principle, we grew populations of non-motile Escherichia coli and motile Pseudomonas aeruginosa for 60 hours with continuous propagation of the colonys front, in contrast to agar plates where growth declined much earlier and stopped after about 40 hours. To demonstrate the capabilities of spatial control, we grew P. aeruginosa along different temporally-stable gradients of the cephalosporin antibiotic ceftazidime and characterised the emerging bacterial growth patterns. The device is a step towards highly controlled studies of microbial populations in continuous, non-uniform spatially structured environments. Designed with cost and accessibility in mind, we believe that this novel device will enable new insights into microbial ecology and evolution.

High-throughput droplet microfluidics can compartmentalize bacterial interactions, but recovering droplets displaying phenotypes of interest often requires custom fluorescence-activated droplet-sorting instrumentation. Here, we introduce post-assay photogelation to decouple the material requirements of bacterial coculture from those of commercial flow sorting. Bacteria are cocultured in initially aqueous water-in-oil droplets containing photoreactive polymer precursors. After interaction phenotypes develop, ultraviolet exposure converts the droplets into mechanically stable hydrogel particles that can be transferred to an aqueous carrier and sorted using a commercial benchtop cell sorter. The sorted particles can subsequently be degraded enzymatically to release the encapsulated bacteria. We show that the timing of gelation alters bacterial growth and spatial distribution within droplets, with post-assay gelation supporting greater and more uniformly distributed growth than culture in preformed hydrogels. Using two fluorescent bead-encoded hydrogel-particle populations, we demonstrate sorting to greater than 99% purity. As an end-to-end demonstration, we cocultured sfGFP-expressing Escherichia coli Nissle 1917 with a cultured human nasal bacterial community and found that E. coli Nissle became the predominant detectable population under the tested conditions with possible inhibition of the cultured nasal bacteriome. This liquid-to-solid transition provides an accessible interface between aqueous bacterial droplet assays, commercial particle sorting, and downstream microbial analysis.

Kidney-function assessment relies on blood urea as a clinically informative metabolic marker; however, its dependence on venipuncture and centralised laboratory testing limits high-frequency monitoring and delays timely clinical intervention. Here, we report an integrated platform combining a wearable buffered microfluidic patch with a physiology-informed, data-driven calibration framework for real-time, non-invasive estimation of blood urea from microlitre-scale sweat volumes (4.79 L). By precisely regulating the release kinetics of internal buffer salts, the device stabilises the local reaction microenvironment, mitigating variability in sweat pH and flow to ensure reproducible measurement. The resulting signals are processed through an artificial intelligence (AI)-enabled analysis pipeline that integrates sweat urea with patient-specific physiological information to generate clinically interpretable outputs. In multicentre studies, sweat urea shows a strong association with blood urea across diverse cohorts, but with nonlinear and time-lagged relationships that limit direct use. The AI-enabled calibration model compensates for these effects, enabling high-fidelity estimation of blood urea (r = 0.945 versus gold-standard measurements) at clinically relevant concordance levels. The platform further identifies kidney injury with 89.1% accuracy and stratifies disease severity with 83.2% accuracy. Notably, these results demonstrate that the integration of physicochemical stabilisation and AI-enabled data-driven translation establishes sweat as a clinically actionable surrogate for renal monitoring, supporting population-level estimation and highlighting the potential for personalised longitudinal assessment, and enabling a scalable, non-invasive strategy for high-frequency kidney disease management.

Perfusable vascular microphysiological systems are increasingly used to model angiogenesis, tissue crosstalk, and disease. However, many platforms still rely on oscillatory, discontinuous, or poorly controlled perfusion regimes, limiting the study of sustained flow-dependent vascular remodeling. Here, we establish a tunable, unidirectional laminar flow workflow for long-term perfusion of angiogenic vasculature-on-chip cultures and use it to investigate endothelial, perivascular, and immune cell responses to sustained flow. Using an AIM Biotech microfluidic platform containing 14-day-old human umbilical vein endothelial cell-derived angiogenic sprouts and pericytes, continuous perfusion enabled intraluminal transport of 1 m tracer beads through vessels, demonstrating stable flow across the vascular bed. Sustained laminar flow induced endothelial remodeling at both the mother vessel and sprout levels, with cellular alignment evident in both compartments. Quantitative analysis of the mother vessel further revealed Golgi polarization against the direction of flow. Sustained perfusion also increased pericyte recruitment to angiogenic sprouts and reduced endothelial proliferation within the mother vessel, consistent with flow-driven vascular maturation and quiescence. Live-cell imaging further captured directional endothelial migration against the flow, lumen remodeling, and dynamic pericyte behavior under continuous perfusion. In immune-cell assays performed under continuous-flow conditions, interactions with untreated endothelium were limited, whereas inflammatory activation increased immune-cell adhesion and crawling. These observations suggest that sustained flow supports a quiescent endothelial phenotype and demonstrate the suitability of the platform for studying inflammatory activation and immune-vascular communication under controlled hemodynamic conditions. Beyond its biological relevance, the workflow provides practical advantages for live-cell imaging, low medium consumption, and downstream perturbation studies. Moreover, the modular design of the platform makes it well suited for vascular-organ crosstalk applications. Collectively, these results establish laminar flow angiogenic vasculature-on-chip as an experimentally tractable model for studying vascular mechanobiology, vascular maturation, and dynamic cell interactions under defined hemodynamic conditions.

Electrophysiological interfacing remains difficult in three-dimensional in vitro models, when using planar microelectrode arrays or optical methods. This challenge limits experimental progress using such models, despite their promise of better physiological relevance than monolayer cell culture. Mesh MEAs which can integrate conformally on or even within tissue offer a possible solution but are not yet widely accessible. Here, we present a mesh MEA device, designed as a simple, manufacturable platform for spheroid electrophysiology. In neural spheroids, the device enabled longitudinal electrophysiological recordings and pharmacological modulation of spontaneous electrical activity. On native polyimide meshes, spheroids maintained their shape while cells enveloped the mesh, embedding electrodes to a depth of 100 {micro}m after 2 weeks. In contrast, laminin biofunctionalization of the mesh promoted outgrowth and migration of cells. This device and associated methods should be adaptable to organoids, ex vivo tissue, or bioengineered in vitro models.

Automation of organoid and cell culture processes is essential for achieving scalable and standardized experimentation in regenerative medicine and stem cell research. However, existing microfluidic platforms often rely on complex setups, limiting their integration within standard incubator environments. To address these challenges, we developed a compact, scalable multi-well platform featuring 3D-printed, servo-actuated disposable microvalves for fully automated media and drug exchange. This design eliminates the need for external pressure sources and control channels, providing a simplified and cost-effective solution for organoid culture. The platform integrates an internet-connected microscopy module with a motorized XYZ stage, allowing continuous, real-time imaging of individual wells directly within the incubator. It supports precise and reliable fluid handling under physiological conditions, improving throughput, reproducibility, and accessibility. We validate the platform through bench-top testing and in both mouse and human organoid models. Morphological analysis, immunohistochemistry (IHC), and qPCR demonstrate comparable viability, growth, and gene expression profiles between automated and manual culture conditions. These results establish a robust and scalable framework for fully automated organoid culture, offering a simplified and accessible alternative to conventional microfluidic systems with broad applications in regenerative medicine, drug discovery, and scalable biological screening. O_FIG O_LINKSMALLFIG WIDTH=200 HEIGHT=101 SRC="FIGDIR/small/732526v1_ufig1.gif" ALT="Figure 1"> View larger version (42K): org.highwire.dtl.DTLVardef@5efd07org.highwire.dtl.DTLVardef@3600d0org.highwire.dtl.DTLVardef@16f85f5org.highwire.dtl.DTLVardef@c39fbd_HPS_FORMAT_FIGEXP M_FIG C_FIG

Adoptive T-cell therapies rely on the identification and expansion of rare tumor-reactive T cells, yet current enrichment strategies are limited by the low abundance of these cells and complexity of their functional enrichment. Here, we present a microfluidic platform that exploits hydrodynamic shear as a controllable parameter for enriching antigen-specific T cells through peptide-major histocompatibility complex (pMHC)-mediated capture. An eight-channel microfluidic device was engineered to simultaneously interrogate a range of wall shear stresses while maintaining uniform cell delivery, enabling systematic identification of shear conditions that maximize antigen-specific enrichment. Using engineered MART-1-specific Jurkat cells, we demonstrate that T-cell capture is jointly regulated by wall shear stress and pMHC density, with intermediate shear preferentially enriching antigen-specific cells over nonspecific binders. Translation of the optimal operating condition to a high-throughput single-shear device enabled approximately 35-fold enrichment of antigen-specific T cells from peripheral blood mononuclear cells containing only 0.05% target cells. We further show that peptide-MHC complexes isolated directly from melanoma whole-cell lysates support shear-dependent enrichment comparable to recombinant pMHCs. Finally, primary MART-1-specific CD8 T cells enriched using tumor-derived pMHCs retained the ability to recognize melanoma cells and upregulated the activation marker CD137 following antigen-specific stimulation. Together, these findings establish hydrodynamic shear as an orthogonal parameter for antigen-specific T-cell enrichment and provide a framework for integrating force-based selection with tumor-derived pMHCs to isolate functional antigen-specific T cells using tumor-derived pMHCs.

Treatment with chimeric antigen receptor (CAR) T cells has emerged as a promising immune therapy for relapsed and refractory hematologic malignancies. The CAR T cells are manufactured in a series of steps that involve isolating T cells from the patients leukapheresis product, genetically modifying them to express the CAR against the target antigen, and reinfusing them into the patient. Efficient T-cell enrichment from leukapheresis products is critical to the success of these therapies. Current methods for T-cell sorting on a clinical scale involve several washing steps to remove red blood cells and platelets, followed by T-cell selection and activation. These multi-step processes result in cell loss during processing and involve several handling steps. Here, we utilize fluidically assembled micromagnetic lenses to develop a high-throughput, continuous-flow microfluidic T-cell sorter, designated as the T-Chip, for sorting magnetic bead-labeled CD3+ T cells in a single step. Our approach allows direct sorting of T cells in expansion media from leukopaks without any washing steps, effectively removing 99.999% of RBCs and platelets from the leukapheresis product. A single 1-inch x 3-inch T-Chip can process leukapheresis product at a throughput of 60 mL/hr and 2.56 {+/-} 0.12 billion cells/hr. Using this optimized workflow, we demonstrate clinical-scale enrichment of highly pure CD3+ T cells (97.7 {+/-} 1.3%) with high viability (97.0 {+/-} 1.1%) and recovery (87.3 {+/-} 14.8%) in a functionally closed manner. Downstream processing of T cells isolated using the T-Chip yielded potent anti-mesothelin CAR T cells with demonstrated anti-tumor efficacy. Overall, by exploiting precisely engineered magnetic forces and laminar flow, the microfluidic T-Chip overcomes bottlenecks caused by low throughput and enables single-step large-scale T-cell purification for the rapid development of CAR T cells.

ObjectiveStructural adaptations of capillary networks, through angiogenesis, arterialization, and regression, are implicated in many diseases, and gaining a better understanding of the cell-cell interactions that underpin these adaptations may lead to novel therapeutic discoveries for disease management. Endothelial cells and pericytes are the two cell types that comprise capillary networks. Experimental model systems have been developed to study the dynamic interactions between endothelial cells and pericytes, providing valuable insights into capillary development, cell-to-cell communication, and responses to growth factors and therapeutic agents. MethodsIn this study, we present a novel and simple co-culture system that uses commercially available primary human endothelial cells and pericytes, does not require microfluidic perfusion, and allows simultaneous observation of cell morphologies and interactions over time in 60 samples, enabling high-throughput analysis of multiple culture conditions with replicates. ResultsImage analysis pipelines were created to quantify microvascular adaptations, including one to measure colocalization between endothelial cells and pericytes, capturing dynamic coupling and uncoupling associated with capillary stability, angiogenesis, and regression. We validated the ability of our co-culture system to reproducibly represent the effects of fibrotic and angiogenic activation signals, including an FDA-approved drug, on endothelial cells, pericytes, and their coupling. ConclusionThis novel, high-throughput microvascular screening assay enables quantification of microvascular dynamics in response to disease-relevant stimuli and therapeutics in a repeatable, real-time manner.

Development of candidate countermeasures against human pathogens frequently includes nonhuman animal experimentation. Preclinical animal pathogen exposure studies are conducted to model diseases and accumulate preliminary and hypothetically translatable data to inform and justify the design of clinical trial evaluation of countermeasure safety and efficacy. In addition to frequent ethical critiques, challenges associated with animal experimentation include considerable resources needed to achieve statistical power and robustness, replicability and reproducibility concerns, potentially compromised objectivity through lack of blinding, fundamental species-specific biological differences, and risk of unpredictable pathogen adaptation to the experimental animal. Recent U.S. and U.K. government initiatives aim to reduce animal experimentation by complementing or potentially replacing them with new approach methodologies (NAMs), i.e., increasingly sophisticated in silico, in chemico, and in vitro approaches. We piloted development of one type of NAM, organ-on-chips (OOCs), in the highly challenging environment of a maximum (biosafety level 4) containment laboratory. Using a Risk Group 4 virus, Nipah virus (NiV), and two types of lung OOCs seeded with human or porcine cells, we demonstrated the recapitulation of key features of NiV lung infection, including viral infection, replication, and translocation, that are associated with proinflammatory cytokine secretion, immune cell recruitment, and disruption of the air-liquid interface barrier. We reproduced the known anti-NiV activity of remdesivir and evaluated that of another potential antiviral, zotatifin. Our results pave the way for similar applications of advanced microphysiological systems for modeling infections caused by high-consequence viruses.