No abstract

The concept of high liquid repellency in multi-liquid-phase systems (e.g., aqueous droplets in an oil background) has been applied to areas of biomedical research to realize intrinsic advantages not available in single-liquid-phase systems. Such advantages have included minimizing analyte loss, facile manipulation of single-cell samples, elimination of biofouling, and ease of use regarding loading and retrieving of the sample. In this paper, we present generalized design rules for predicting the wettability of solid-liquid-liquid systems (especially for discrimination between exclusive liquid repellency (ELR) and finite liquid repellency) to extend the applications of ELR. We then apply ELR to two model systems with open microfluidic design in cell biology: (1) in situ underoil culture and combinatorial coculture of mammalian cells in order to demonstrate directed single-cell multiencapsulation with minimal waste of samples as compared to stochastic cell seeding and (2) isolation of a pure population of circulating tumor cells, which is required for certain downstream analyses including sequencing and gene expression profiling.

Double-exclusive liquid repellency (double-ELR) is an extreme wettability phenomenon in which adjacent regions selectively and completely repel immiscible liquids with different surface chemistries on a non-textured substrate (i.e., a substrate in absence of micro/nano-structures). Under double-ELR conditions, each liquid exhibits no physical contact (contact angle of 180°) with its non-preferred surface chemistry, thus enabling complete partitioning of adjacent fluidic volumes (e.g., between water and oil). This enables a new type of cell culture-based assay, where cell loss from common failure modes (e.g., biofouling from inadvertent cell adhesion, detrimental moisture loss/gain, and liquid handling dead volumes) is significantly mitigated. Importantly, the principles of double-ELR were leveraged to achieve underoil sweep patterning, a no-loss, robust and high-throughput distribution of sub-microliter volumes of aqueous media (and cells). In addition to high-efficiency distribution via sweep patterning, double-ELR can be used to construct "modular" (i.e., easily implemented and/or linked together with spatial and temporal control) higher-order architectures for in vitro imitation of physiologically relevant microenvironments that are of particular interest within the cell assay community, including multi-phenotype cultures with excellent spatial and temporal control, three-dimensional layered multi-phenotype cultures, cultures with selective mechanical cues of extracellular matrix (i.e., collagen fiber alignment), and spheroid cultures. Together, these features of double-ELR uniquely facilitate culture and high content analysis of limited cellular samples (e.g., a few hundred to a few thousand cells).

Angiogenesis (the formation of blood vessels from existing blood vessels) plays a critical role in many diseases such as cancer, benign tumors, and macular degeneration. There is a need for cell culture methods capable of dissecting the intricate regulation of angiogenesis within the microenvironment of the vasculature. We have developed a microscale cell-based assay that responds to complex pro- and anti-angiogenic soluble factors with an in vitro readout for vessel formation. The power of this system over traditional techniques is that we can incorporate the whole milieu of soluble factors produced by cells in situ into one biological readout (vessel formation), even if the identity of the factors is unknown. We have currently incorporated macrophages, endothelial cells, and fibroblasts into the assay, with the potential to include additional cell types in the future. Importantly, the microfluidic platform is simple to operate and multiplex to test drugs targeting angiogenesis in a more physiologically relevant context. As a proof of concept, we tested the effect of an enzyme inhibitor (targeting matrix metalloproteinase 12) on vessel formation; the triculture microfluidic assay enabled us to capture a dose-dependent effect entirely missed in a simplified coculture assay (p<0.0001). This result underscores the importance of cell-based assays that capture chemical cross-talk occurring between cell types. The microscale dimensions significantly reduce cell consumption compared to conventional well plate platforms, enabling the use of limited primary cells from patients in future investigations and offering the potential to screen therapeutic approaches for individual patients in vitro.

Micromilling is an underutilized technique for fabricating microfluidic platforms that is well-suited for the diverse needs of the biologic community. This technique, however, produces culture surfaces that are considerably rougher than in commercially available culture platforms and the hydrophilicity of these surfaces can vary considerably depending on the choice of material. In this study, we evaluated the impact of surface topography and hydrophilicity in milled microfluidic devices on the cellular phenotype and function of primary human macrophages. We found that the rough culture surface within micromilled systems affected the phenotype of macrophages cultured in these devices. However, the presence, type, and magnitude of this effect was dependent on the surface hydrophilicity as well as exposure to chemical polarization signals. These findings confirm that while milled microfluidic systems are an effective platform for culture and analysis of primary macrophages, the topography and hydrophilicity of the culture surface within these systems should be considered in the planning and analysis of any macrophage experiments in which phenotype is relevant.

Summary Mitotic arrest deficient 1 (Mad1) plays a well-characterized role in the major cell cycle checkpoint that regulates chromosome segregation during mitosis, the mitotic checkpoint (also known as the spindle assembly checkpoint). During mitosis, Mad1 recruits Mad2 to unattached kinetochores [1, 2], where Mad2 is converted into an inhibitor of the anaphase promoting complex/cyclosome bound to its specificity factor Cdc20 [1, 3–6]. During interphase, Mad1 remains tightly bound to Mad2 [2, 3, 7, 8] and both proteins localize to the nucleus and nuclear pores [9, 10], where they interact with Tpr (Translocated Promoter Region). Recently, it has been shown that interaction with Tpr stabilizes both proteins [11], and that Mad1 binding to Tpr permits Mad2 to associate with Cdc20 [12]. However, interphase functions of Mad1 that do not directly affect the mitotic checkpoint have remained largely undefined. Here we identify a previously unrecognized interphase distribution of Mad1 at the Golgi apparatus. Mad1 colocalizes with multiple Golgi markers and cosediments with Golgi membranes. Although Mad1 has previously been thought to constitutively bind Mad2, Golgi-associated Mad1 is Mad2-independent. Depletion of Mad1 impairs secretion of α5 integrin and results in defects in cellular attachment, adhesion, and FAK activation. Additionally, reduction of Mad1 impedes cell motility, while its overexpression accelerates directed cell migration. These results reveal an unexpected role for a mitotic checkpoint protein in secretion, adhesion and motility. More generally, they demonstrate that, in addition to generating aneuploidy, manipulation of mitotic checkpoint genes can have unexpected interphase effects that influence tumor phenotypes.

Macrophages within the tumor microenvironment (TME) exhibit a spectrum of pro-tumor and antitumor functions, yet it is unclear how the TME regulates this macrophage heterogeneity. Standard methods to measure macrophage heterogeneity require destructive processing, limiting spatiotemporal studies of function within the live, intact 3D TME. Here, we demonstrate twophoton autofluorescence imaging of NAD(P)H and FAD to non-destructively resolve spatiotemporal metabolic heterogeneity of individual macrophages within 3D microscale TME models. Fluorescence lifetimes and intensities of NAD(P)H and FAD were acquired at 24, 48, and 72 hours post-stimulation for mouse macrophages (RAW 264.7) stimulated with IFN-γ or IL-4 plus IL-13 in 2D culture, validating that autofluorescence measurements capture known metabolic phenotypes. To quantify metabolic dynamics of macrophages within the TME, mouse macrophages or human monocytes (RAW264.7 or THP-1) were cultured alone or with breast cancer cells (mouse PyVMT or primary human IDC) in 3D microfluidic platforms. Human monocytes and mouse macrophages in tumor co-cultures exhibited significantly different FAD mean lifetimes and greater migration than mono-cultures at 24, 48, and 72 hours post-seeding. In co-cultures with primary human cancer cells, actively-migrating monocyte-derived macrophages had greater redox ratios (NAD(P)H/FAD intensity) compared to passively-migrating monocytes at 24 and 48 hours post-seeding, reflecting metabolic heterogeneity in this subpopulation of monocytes. Genetic analyses further confirmed this metabolic heterogeneity. These results establish label-free autofluorescence imaging to quantify dynamic metabolism, polarization, and migration of macrophages at single-cell resolution within 3D microscale models. This combined culture and imaging system provides unique insights into spatiotemporal tumorimmune crosstalk within the 3D TME.

Macrophages within the tumor microenvironment (TME) exhibit a spectrum of pro-tumor and anti-tumor functions, yet it is unclear how the TME regulates this macrophage heterogeneity. Standard methods to measure macrophage heterogeneity require destructive processing, limiting spatiotemporal studies of function within the live, intact 3D TME. Here we demonstrate two-photon autofluorescence imaging of NAD(P)H and FAD to non-destructively resolve spatiotemporal metabolic heterogeneity of individual macrophages within 3D microscale TME models. Fluorescence lifetimes and intensities of NAD(P)H and FAD were acquired at 24, 48, and 72 hours post-stimulation for mouse macrophages (RAW 264.7) stimulated with IFN-γ or IL-4 plus IL-13 in 2D culture, confirming that autofluorescence measurements capture known metabolic phenotypes. To quantify metabolic dynamics of macrophages within the TME, mouse macrophages or human monocytes (RAW264.7 or THP-1) were cultured alone or with breast cancer cells (mouse PyVMT or primary human IDC) in 3D microfluidic platforms. Human monocytes and mouse macrophages in tumor co-cultures exhibited significantly different FAD mean lifetimes and greater migration than monocultures at 24, 48, and 72 hours post-seeding. In co-cultures with primary human cancer cells, actively-migrating monocyte-derived macrophages had greater redox ratios (NAD(P)H/FAD intensity) compared to passivelymigrating monocytes at 24 and 48 hours post-seeding, reflecting metabolic heterogeneity in this sub-population of monocytes. Genetic analyses further confirmed this metabolic heterogeneity. These results establish label-free autofluorescence imaging to quantify dynamic metabolism, polarization, and migration of macrophages at single-cell resolution within 3D microscale models. This combined culture and imaging system provides unique insights into spatiotemporal tumor-immune crosstalk within the 3D TME. SIGNIFICANCE Label-free metabolic imaging and microscale culture technologies enable monitoring of singlecell macrophage metabolism, migration, and function in the 3D tumor microenvironment. Research.