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Vascular hypo‐fibrinolysis is a historically underappreciated and understudied aspect of venous thromboembolism (VTE). This paper describes the development of a micro‐clot dissolution assay for quantifying the fibrinolytic capacity of endothelial cells – a key driver of VTE development. This assay is enabled using aqueous two‐phase systems (ATPS) to bioprint microscale fibrin clots over human umbilical vein endothelial cells (HUVECs). Importantly, these micro‐clots are orders of magnitude smaller than conventional fibrin constructs and allow HUVEC‐produced plasminogen activators to mediate visually quantifiable fibrinolysis. Using live‐cell time‐lapse imaging, micro‐clot dissolution by HUVECs is tracked, and fibrinolysis kinetics are quantified. The sensitivity of cell‐driven fibrinolysis to various stimuli is rapidly tested. The physiological relevance of this convenient high‐throughput assay is illustrated through treatments with lipopolysaccharide (LPS) and rosuvastatin that elicit anti‐ and pro‐fibrinolytic responses, respectively. Furthermore, treatment with baricitinib, an anti‐inflammatory therapeutic found to increase cardiovascular risks after market approval, provokes an anti‐fibrinolytic response – which highlights the potential role of endothelial cells in increasing VTE risk for patients receiving this drug. This endothelial cell fibrinolysis assay provides a high‐throughput and versatile drug testing platform – potentially allowing for early preclinical identification of therapeutics that may beneficially enhance or adversely impair endothelial fibrinolysis.

Live-cell imaging reveals the phenotypes and mechanisms of cellular function and their dysfunction that underscore cell physiology, development, and pathology. Here, we report a 3D super-resolution live-cell microscopy method by integrating radiality analysis and Fourier light-field microscopy (rad-FLFM). We demonstrated the method using various live-cell specimens, including actins in Hela cells, microtubules in mammary organoid cells, and peroxisomes in COS-7 cells. Compared with conventional wide-field microscopy,rad-FLFM realizes scanning-free, volumetric 3D live-cell imaging with sub-diffraction-limited resolution of ∼150 nm (x-y) and 300 nm (z), milliseconds volume acquisition time, six-fold extended depth of focus of ∼6 µm, and low photodamage. The method provides a promising avenue to explore spatiotemporal-challenging subcellular processes in a wide range of cell biological research.

This paper describes a microscale fibroplasia and contraction model that is based on fibrin-embedded lung fibroblasts and provides a convenient visual readout of fibrosis. Cell-laden fibrin microgel drops are formed by aqueous two-phase microprinting. The cells deposit extracellular matrix (ECM) molecules such as collagen while fibrin is gradually degraded. Ultimately, the cells contract the collagen-rich matrix to form a compact cell-ECM spheroid. The size of the spheroid provides the visual readout of the extent of fibroplasia. Stimulation of this wound-healing model with the profibrotic cytokine TGF-β1 leads to an excessive scar formation response that manifests as increased collagen production and larger cell-ECM spheroids. Addition of drugs also shifted the scarring profile: the FDA-approved fibrosis drugs (nintedanib and pirfenidone) and a PAI-1 inhibitor (TM5275) significantly reduced cell-ECM spheroid size. Not only is the assay useful for evaluation of antifibrotic drug effects, it is relatively sensitive; one of the few in vitro fibroplasia assays that can detect pirfenidone effects at submillimolar concentrations. Although this paper focuses on lung fibrosis, the approach opens opportunities for studying a broad range of fibrotic diseases and for evaluating antifibrotic therapeutics.

3D in vitro model systems such as spheroids and organoids provide an opportunity to extend the physiological understanding using recapitulated tissues that mimic physiological characteristics of in vivo microenvironments. Unlike 2D systems, 3D in vitro systems can bridge the gap between inadequate 2D cultures and the in vivo environments, providing novel insights on complex physiological mechanisms at various scales of organization, ranging from the cellular, tissue‐, to organ‐levels. To satisfy the ever‐increasing need for highly complex and sophisticated systems, many 3D in vitro models with advanced microengineering techniques have been developed to answer diverse physiological questions. This review summarizes recent advances in engineered microsystems for the development of 3D in vitro model systems. The relationship between the underlying physics behind the microengineering techniques, and their ability to recapitulate distinct 3D cellular structures and functions of diverse types of tissues and organs are highlighted and discussed in detail. A number of 3D in vitro models and their engineering principles are also introduced. Finally, current limitations are summarized, and perspectives for future directions in guiding the development of 3D in vitro model systems using microengineering techniques are provided.

High‐throughput tissue barrier models can yield critical insights on how barrier function responds to therapeutics, pathogens, and toxins. However, such models often emphasize multiplexing capability at the expense of physiologic relevance. Particularly, the distal lung's air–blood barrier is typically modeled with epithelial cell monoculture, neglecting the substantial contribution of endothelial cell feedback in the coordination of barrier function. An obstacle to establishing high‐throughput coculture models relevant to the epithelium/endothelium interface is the requirement for underside cell seeding, which is difficult to miniaturize and automate. Therefore, this paper describes a scalable, low‐cost seeding method that eliminates inversion by optimizing medium density to float cells so they attach under the membrane. This method generates a 96‐well model of the distal lung epithelium–endothelium barrier with serum‐free, glucocorticoid‐free air–liquid differentiation. The polarized epithelial–endothelial coculture exhibits mature barrier function, appropriate intercellular junction staining, and epithelial‐to‐endothelial transmission of inflammatory stimuli such as polyinosine:polycytidylic acid (poly(I:C)). Further, exposure to influenza A virus PR8 and human beta‐coronavirus OC43 initiates a dose‐dependent inflammatory response that propagates from the epithelium to endothelium. While this model focuses on the air–blood barrier, the underside seeding method is generalizable to various coculture tissue models for scalable, physiologic screening.

Extracellular traps (ETs), such as neutrophil extracellular traps, are a physical mesh deployed by immune cells to entrap and constrain pathogens. ETs are immunogenic structures composed of DNA, histones, and an array of variable protein and peptide components. While much attention has been paid to the multifaceted function of these structures, mechanistic studies of ETs remain challenging due to their heterogeneity and complexity. Here, a novel DNA‐histone mesostructure (DHM) formed by complexation of DNA and histones into a fibrous mesh is reported. DHMs mirror the DNA‐histone structural frame of ETs and offer a facile platform for cell culture studies. It is shown that DHMs are potent activators of dendritic cells and identify both the methylation state of DHMs and physical interaction between dendritic cells and DHMs as key tuning switches for immune stimulation. Overall, the DHM platform provides a new opportunity to study the role of ETs in immune activation and pathophysiology.