Mario Matteo Modena

Three-dimensional (3D) cell models, such as spheroids and organoids, are powerful tools to enhance the predictive power of preclinical in vitro studies and support clinical translation of anti-cancer therapies. However, their integration into drug discovery pipelines remains limited, particularly for high-throughput screening, where rapid and reliable viability assessment is essential. Conventional viability assays, often dye-based or destructive, remain labor-intensive and poorly suited for high-throughput applications and downstream re-use of the models. In this study, we demonstrate, for the first time, that electrical impedance spectroscopy (EIS) enables label-free viability profiling of individual spheroids. For this purpose, a differential EIS sensor was integrated into a plastic microfluidic chip in flow-through configuration with self-aligned facing electrodes. Using six probing frequencies, we measured the EIS responses of liver cancer spheroids exposed to varying concentrations of dimethyl sulfoxide and doxorubicin, a chemotherapeutic compound. We identified an optimized frequency pair that enabled us to monitor spheroid viability, independent of spheroid size. Comparison with conventional viability assays, namely fluorescence imaging and ATP quantification, confirmed that the EIS method reliably reflected viability changes and revealed inter- and intra-treatment group differences owing to the method’s single-spheroid resolution. The same frequency pair could be used to detect viability changes for both compounds, suggesting transferability of the EIS method across treatments. Furthermore, we showed that utilizing additional EIS features improved classification of spheroids with subtle viability differences. Together, these results demonstrate that EIS is a promising approach for noninvasive, single-spheroid viability profiling and a valuable tool for high-throughput drug screening with spheroid models.

Organotypic brain slice cultures (OSCs) provide a physiologically relevant model for recapitulating complex tissues in vitro. However, the longitudinal monitoring of dynamic processes in OSCs through high-resolution imaging is limited in standard culturing methods, as they rely on slice placement on porous membranes at the air-liquid interface to preserve tissue viability. Here, a new experimental paradigm based on a modular slice culture insert is introduced, which is compatible with static and perfused culturing, and facilitates longitudinal observation of cellular and tissue dynamics at high spatiotemporal resolution. Besides compatibility with conventional culturing in well plates, the insert is integrated into an automated microfluidic instrument to provide user-friendly operation while enabling precise temporal fluid control and compound delivery. The system is applied to monitor calcium responses and cellular morphology of virally labeled cerebellar astrocytes and Purkinje cells exposed to different concentrations of glutamate. Cellular responses were evaluated at baseline, during glutamate exposure, and 48 h and 96 h after stimulation to identify acute and long-term excitotoxic effects. Our results indicate that the developed system enables high-resolution monitoring of cellular morphology and dynamic events of the same intact tissue over multiple days, which provides a robust foundation for mechanistic investigations of neural signaling.

Few therapeutic options for treating neurological diseases are currently available due to the extremely selective nature of the blood-brain barrier (BBB), which strongly limits drug delivery from the systemic circulation into the central nervous system. The lack of effective treatment options is particularly dire for brain tumors, which ultimately result in very low survival rates. To address the challenge of evaluating drug permeation and efficacy within a physiologically relevant context, we developed a perfused, open-microfluidic platform that includes a human BBB model in co-culture with tumor spheroids. The platform was fabricated from inert plastics to enable quantitative small molecule testing, and it featured 32 testing units in a well-plate format. A pump-free, gravity-driven flow scheme was adopted to establish physiological shear-stress conditions and to enable simple parallelization on tilting stages for increased throughput. We tested the efficacy and permeation of four FDA-approved small-molecule chemotherapeutics - cisplatin, doxorubicin, homoharringtonine, and docetaxel on two patient-derived diffuse-midline-glioma models at sub-IC₅₀ drug concentrations for the BBB. Our results demonstrate that the in vitro BBB significantly limited drug delivery to the tumor, thereby limiting drug efficacy. Furthermore, drug-induced BBB disruption occurred at sub-toxic doses, which led to increased drug permeation to the glioma models. Finally, cell-model-specific responses revealed distinct cytotoxicity behavior, demonstrating the importance of personalized therapy testing. Our scalable BBB-tumor platform provides a physiologically relevant in vitro model system to assess drug permeation, cytotoxicity, and tumor-BBB interactions and offers the potential to advance the discovery of new effective therapeutics against neurological diseases.

In vitro evaluation of novel therapeutic approaches often fails to reliably predict efficacy and toxicity, especially when recapitulating conditions involving recirculating cells. Current testing strategies are often based on static co-culturing of cells in suspension and 3D tissue models, where cell sedimentation on the target tissue can occur. The observed effects may then mostly be a consequence of sedimentation and of the corresponding forced cell-tissue interactions. The realization of continuous medium flow helps to better recapitulate physiological conditions and cell-tissue interactions. To tackle current limitations of perfused organ-on-chip approaches, we developed a microfluidic chip and operation concept, which prevents undesired sedimentation and accumulation of suspended cells during multiple days by relying on gravity-driven perfusion. Our platform, which we termed “human immune flow (hiFlow) chip”, enables to co-culture cells in suspension with up to 7 preformed microtissue models. Here, we present the design principle and operation of the platform, and we validate its performance by culturing cells and microtissues of a variety of different origins. Cells and tissues could be monitored on chip via high-resolution microscopy, while cell suspensions and microtissues could be easily retrieved for off-chip analysis. Our results demonstrate that primary immune cells and a range of different spheroid models of healthy and diseased tissues can be maintained for over 6 days on chip. As proof-of-concept cell-tissue interaction assay, we used an antibody treatment against diffuse midline glioma, a highly aggressive pediatric tumor. We are confident that our platform will help to increase the prediction power of in vitro preclinical testing of novel therapeutics that rely on the interaction of circulating cells with organ tissues.

As traditional two-dimensional (2D) cell cultures offer limited predictive capabilities for drug development, three-dimensional (3D) tissue models, such as spherical microtissues, have been introduced to better reproduce physiological conditions. The hanging-drop method, used to cultivate microtissues at an air–liquid interface, proves to be effective for microtissue formation and maintenance. Using that technology, it is possible to fluidically interconnect several hanging drops hosting different models of human organs to recapitulate relevant tissue interactions. Here, we combine microfluidics with microelectronics (i.e., complementary metal-oxide-semiconductor (CMOS) technology) and present a novel multifunctional CMOS microelectrode array (MEA) integrated into an open microfluidic system. The device can be used in hanging-drop mode for in situ microtissue readouts and in standing-drop mode like a conventional MEA. The CMOS-MEA chip features two reconfigurable electrode arrays with 1024 electrodes each, and enables electrophysiology, impedance spectroscopy, and electrochemical sensing to acquire a broad spectrum of biologically relevant information. We fabricated the chip using a 0.18 μm CMOS process and developed a strategy to integrate the CMOS-MEA chip into the open microfluidic system within a larger overall effort to incorporate discrete CMOS sensors into microfluidic devices. Proof-of-concept experiments demonstrate the capability to perform electrophysiology and impedance spectroscopy of human induced pluripotent stem cell (hiPSC)-derived cardiac microtissues, as well as electrochemical sensing of different analytes including hydrogen peroxide and epinephrine.

Replicating the cytoarchitecture and cell heterogeneity of the brain in vitro remains challenging. Although ex vivo organotypic slices preserve native tissue complexity, current culturing methods limit long-term, high-resolution imaging and restrict temporally controlled perturbations. Here, we present PHIROS, a microfluidic platform for high-resolution imaging of organotypic slices that supports static culturing of excised tissue at the air–liquid interface and provides continuous optical access for extended imaging at subcellular resolution. Controlled perfusion with oxygenated medium preserves tissue viability over several days, enabling, e.g., the monitoring of spontaneous and pharmacologically modulated astrocytic calcium activity. Using PHIROS we characterized medulloblastoma (MB) cell behavior in a physiological tumor microenvironment and observed dynamic F-actin-driven interactions with tissue-resident astrocytes, as well as leading edge localization of the immune-checkpoint marker B7-H3 in invading tumor cells. Quantitative assessment of mitochondria transfer across heterotypic actin-rich connections evidences the potential of PHIROS as a versatile system for mechanistic studies in a tissue context, enabling controlled compound exposure and high-resolution imaging in physiologically relevant tissue settings.

In vitro evaluation of novel therapeutic approaches often fails to reliably predict efficacy and toxicity, especially when recapitulating conditions involving recirculating cells. Current testing strategies are often based on static co-culturing of cells in suspension and 3D tissue models, where cell sedimentation on the target tissue can occur. The observed effects may then mostly be a consequence of sedimentation and of the corresponding forced cell-tissue interactions. The realization of continuous medium flow helps to better recapitulate physiological conditions. To tackle current limitations of perfused organ-on-chip approaches, we developed a microfluidic chip and operation concept, which prevents undesired sedimentation and accumulation of cells in suspensions over multiple days by relying on gravity-driven perfusion. Our platform, which we termed “human immune flow (hiFlow) chip”, enables to co-culture cells in suspension with up to 7 preformed microtissue models. The primary goal was to study flow-dependent recruitment of the circulating cells to the different tissue spheroids. Cells and tissues could be monitored on-chip via high-resolution microscopy, while cell suspensions and microtissues could be easily retrieved for off-chip analysis. Our results demonstrate that primary immune cells and spheroid models of healthy and diseased tissue can be maintained for over 6 days on chip. As proof-of-principle assay, we investigated the efficacy of antibody treatment against diffuse midline glioma, a highly aggressive pediatric tumor. We believe that our platform will help to increase the prediction power of in vitro preclinical testing of novel immunotherapeutics.

Electrical impedance spectroscopy (EIS) is a promising label-free tool for high-throughput analysis of 3D cellular constructs, also called spheroids. Here, we used an EIS platform featuring facing electrodes to characterize the viability of hepatic spheroids, which are used for bioprinting applications. By using principal component analysis (PCA), we show that this simple impedance sensor enables us to successfully distinguish healthy spheroids from spheroids exposed to toxic conditions. The sensitivity of the impedance sensor will be further characterized by using spheroids exposed to varying stress conditions like different drug concentrations and temperatures.

Blood‐brain‐barrier (BBB) disruption has been associated with a variety of central‐nervous‐system diseases. In vitro BBB models enable to investigate how the barrier reacts to external injury events, commonly referred to as insults. Here, a human‐cell‐based BBB platform with integrated, transparent electrodes to monitor barrier tightness in real time at high resolution is presented. The BBB model includes human cerebral endothelial cells and primary pericytes and astrocytes in a 3D arrangement within a pump‐free, open‐microfluidic platform. With this platform, this study demonstrates that oxygen‐glucose deprivation (OGD), which mimics the characteristics of an ischemic insult, induces a rapid remodeling of the cellular actin structures and subsequent morphological changes in the endothelial cells. High‐resolution live imaging shows the formation of large actin stress‐fiber bundles in the endothelial layer during OGD application, which ultimately leads to cell shrinkage and barrier breakage. Simultaneous electrical measurements evidence a rapid decrease of the barrier electrical resistance before the appearance of stress fibers, which indicates that the barrier function is compromised already before the appearance of drastic morphological changes. The results demonstrate that the BBB platform recapitulates the main barrier functions in vitro and can be used to investigate rapid reorganization of the BBB upon application of external stimuli.

Despite increasing survival rates of pediatric leukemia patients over the past decades, the outcome of some leukemia subtypes has remained dismal. Drug sensitivity and resistance testing on patient‐derived leukemia samples provide important information to tailor treatments for high‐risk patients. However, currently used well‐based drug screening platforms have limitations in predicting the effects of prodrugs, a class of therapeutics that require metabolic activation to become effective. To address this issue, a microphysiological drug‐testing platform is developed that enables co‐culturing of patient‐derived leukemia cells, human bone marrow mesenchymal stromal cells, and human liver microtissues within the same microfluidic platform. This platform also enables to control the physical interaction between the diverse cell types. Herein, it is made possible to recapitulate hepatic prodrug activation of ifosfamide in their platform, which is very difficult in traditional well‐based assays. By testing the susceptibility of primary patient‐derived leukemia samples to the prodrug ifosfamide, sample‐specific sensitivities to ifosfamide in primary leukemia samples are identified. The microfluidic platform is found to enable the recapitulation of physiologically relevant conditions and the testing of prodrugs including short‐lived and unstable metabolites. The platform holds great potential for clinical translation and precision chemotherapy selection.