Nerses J. Haroutunian

Drug development suffers from a lack of predictive and human-relevant in vitro models. Organ-on-chip (OOC) technology provides advanced culture capabilities to generate physiologically appropriate, human-based tissue in vitro, therefore providing a route to a predictive in vitro model. However, OOC technologies are often created at the expense of throughput, industry-standard form factors, and compatibility with state-of-the-art data collection tools. Here we present an OOC platform with advanced culture capabilities supporting a variety of human tissue models including liver, vascular, gastrointestinal, and kidney. The platform has 96 devices per industry standard plate and compatibility with contemporary high-throughput data collection tools. Specifically, we demonstrate programmable flow control over two physiologically relevant flow regimes: perfusion flow that enhances hepatic tissue function and high-shear stress flow that aligns endothelial monolayers. In addition, we integrate electrical sensors, demonstrating quantification of barrier function of primary gut colon tissue in real-time. We utilize optical access to the tissues to directly quantify renal active transport and oxygen consumption via integrated oxygen sensors. Finally, we leverage the compatibility and throughput of the platform to screen all 96 devices using high content screening (HCS) and evaluate gene expression using RNA sequencing (RNA-seq). By combining these capabilities in one platform, physiologically-relevant tissues can be generated and measured, accelerating optimization of an in vitro model, and ultimately increasing predictive accuracy of in vitro drug screening.

Implementation of gene editing technologies such as CRISPR/Cas9 in the manufacture of novel cell-based therapeutics has the potential to enable highly-targeted, stable, and persistent genome modifications without the use of viral vectors. Electroporation has emerged as a preferred method for delivering gene-editing machinery to target cells, but a major challenge remaining is that most commercial electroporation machines are built for research and process development rather than for large-scale, automated cellular therapy manufacturing. Here we present a microfluidic continuous-flow electrotransfection device designed for precise, consistent, and high-throughput genetic modification of target cells in cellular therapy manufacturing applications. We optimized our device for delivery of mRNA into primary human T cells and demonstrated up to 95% transfection efficiency with minimum impact on cell viability and expansion potential. We additionally demonstrated processing of samples comprising up to 500 million T cells at a rate of 20 million cells/min. We anticipate that our device will help to streamline the production of autologous therapies requiring on the order of 10[Formula: see text]-10[Formula: see text] cells, and that it is well-suited to scale for production of trillions of cells to support emerging allogeneic therapies.

Microphysiological organ-on-chip models offer the potential to improve the prediction of drug safety and efficacy through recapitulation of human physiological responses. The importance of including multiple cell types within tissue models has been well documented. However, the study of cell interactions in vitro can be limited by complexity of the tissue model and throughput of current culture systems. Here, we describe the development of a co-culture microvascular model and relevant assays in a high-throughput thermoplastic organ-on-chip platform, PREDICT96. The system consists of 96 arrayed bilayer microfluidic devices containing retinal microvascular endothelial cells and pericytes cultured on opposing sides of a microporous membrane. Compatibility of the PREDICT96 platform with a variety of quantifiable and scalable assays, including macromolecular permeability, image-based screening, Luminex, and qPCR, is demonstrated. In addition, the bilayer design of the devices allows for channel- or cell type-specific readouts, such as cytokine profiles and gene expression. The microvascular model was responsive to perturbations including barrier disruption, inflammatory stimulation, and fluid shear stress, and our results corroborated the improved robustness of co-culture over endothelial mono-cultures. We anticipate the PREDICT96 platform and adapted assays will be suitable for other complex tissues, including applications to disease models and drug discovery.

Hepatic in vitro platforms ranging from multi-well cultures to bioreactors and microscale systems have been developed as tools to recapitulate cellular function and responses to aid in drug screening and disease model development. Recent developments in microfabrication techniques and cellular materials enabled fabrication of next-generation, advanced microphysiological systems (MPSs) that aim to capture the cellular complexity and dynamic nature of the organ presenting highly controlled extracellular cues to cells in a physiologically relevant context. Historically, MPSs have heavily relied on elastomeric materials in their manufacture, with unfavorable material characteristics (such as lack of structural rigidity) limiting their use in high-throughput systems. Herein, we aim to create a microfluidic bilayer model (microfluidic MPS) using thermoplastic materials to allow hepatic cell stabilization and culture, retaining hepatic functional phenotype and capturing cellular interactions. The microfluidic MPS consists of two overlapping microfluidic channels separated by a porous tissue-culture membrane that acts as a surface for cellular attachment and nutrient exchange; and an oxygen permeable material to stabilize and sustain primary human hepatocyte (PHH) culture. Within the microfluidic MPS, PHHs are cultured in the top channel in a collagen sandwich gel format with media exchange accomplished through the bottom channel. We demonstrate PHH culture for 7 days, exhibiting measures of hepatocyte stabilization, secretory and metabolic functions. In addition, the microfluidic MPS dimensions provide a reduced media-to-cell ratio in comparison with multi-well tissue culture systems, minimizing dilution and enabling capture of cellular interactions and responses in a hepatocyte-Kupffer coculture model under an inflammatory stimulus. Utilization of thermoplastic materials in the model and ability to incorporate multiple hepatic cells within the system is our initial step towards the development of a thermoplastic-based high-throughput microfluidic MPS platform for hepatic culture. We envision the platform to find utility in development and interrogation of disease models of the liver, multi-cellular interactions and therapeutic responses.

Abstract Autologous cellular therapies have been highly successful in treating hematological cancers and have the potential to be used for a variety of indications. Manufacturing these therapies rapidly and at low cost remains a major challenge. A key bottleneck in cellular therapy manufacturing is genetic modification of target cells, which is often done using viral vectors. Because vectors are expensive to develop and produce, non‐viral gene transfer using electroporation is emerging as a preferred transfection method for next‐generation therapies. However, most commercial electroporation systems are built for research use rather than large‐scale clinical manufacturing. The microfluidic, continuous‐flow electroporation device presented here offers several advantages including large‐scale and high throughput processing, high performance, and the potential for automation. It transfects primary human T cells with Cas9‐guide ribonucleic acid (RNA) ribonucleoprotein complexes (RNP) and messenger RNA (mRNA) with up to 99–100% efficiency and minimal impact on viability. In addition, this device transfects 3.5 kbp plasmid deoxyribonucleic acid with up to 86% efficiency after preliminary optimization studies. A single microchannel can deliver a total cellular processing throughput of up to 9.6 billion per hour. The combination of high throughput and high performance enables the scale of processing required for future “off‐the‐shelf” allogeneic cellular therapies.