Search

Microfluidic 3D tissue culture systems are attractive for in vitro drug testing applications due to the ability of these platforms to generate 3D tissue models and perform drug testing at a very small scale. However, the minute cell number and liquid volume impose significant technical challenges to perform quantitative cell viability measurements using conventional colorimetric or fluorometric assays, such as MTS or Alamar Blue. Similarly, live-dead staining approaches often utilize metabolic dyes that typically label the cytoplasm of live cells, which makes it difficult to segment and count individual cells in compact 3D tissue cultures. In this paper, we present a quantitative image-based cell viability (QuantICV) assay technique that circumvents current challenges of performing the quantitative cell viability assay in microfluidic 3D tissue cultures. A pair of cell-impermeant nuclear dyes (EthD-1 and DAPI) were used to sequentially label the nuclei of necrotic and total cell populations, respectively. Confocal microscopy and image processing algorithms were employed to visualize and quantify the cell nuclei in the 3D tissue volume. The QuantICV assay was validated and showed good concordance with the conventional bulk MTS assay in static 2D and 3D tumor cell cultures. Finally, the QuantICV assay was employed as an on-chip readout to determine the differential dose responses of parental and metastatic 3D oral squamous cell carcinoma (OSCC) to Gefitinib in a microfluidic 3D culture device. This proposed technique can be useful in microfluidic cell cultures as well as in a situation where conventional cell viability assays are not available.

We report a compartmentalized microbiome module that allows culturing of commensal bacteria under either aerobic or anaerobic conditions, that can be easily connected to our existing modular microfluidic platform. The microbiome module is designed to permit translocation of metabolites across a semi-permeable membrane and circulated to a 3D liver culture module. Our findings showed that microbial metabolites such as sodium butyrate (NaB) can reduce lipid droplet accumulation and pro-inflammation markers in fatty liver cultures.

'Muscle-on-Chip' technologies have been developed to serve as physiologically relevant animal substitutes for in-vitro screening of drugs and study of myogenesis. The formation of 3D suspended myo-bundles within a contractable hydrogel in 'Muscle-on-Chip' devices has been well-established thus far. However, very limited studies have elucidated (i) The effect of physiological shear stresses and (ii) The mechanical properties of the 3D hydrogel microenvironment in myo-bundle formation and their subsequent myogenesis. As such, we have developed a perfusable 'Muscle-on-Chip' device which will enable us to study the effect of these dynamic stimuli on myo-bundle formation and maturation.

Multi-organ perfusion systems offer the unique opportunity to mimic different physiological systemic interactions. However, existing multi-organ culture platforms have limited flexibility in specifying the culture conditions, device architectures, and fluidic connectivity simultaneously. Here, we report a modular microfluidic platform that addresses this limitation by enabling easy conversion of existing microfluidic devices into tissue and fluid control modules with self-aligning magnetic interconnects. This enables a 'stick-n-play' approach to assemble planar perfusion circuits that are amenable to both bioimaging-based and analytical measurements. A myriad of tissue culture and flow control TILE modules were successfully constructed with backward compatibility. Finally, we demonstrate applications in constructing recirculating multi-organ systems to emulate liver-mediated bioactivation of nutraceuticals and prodrugs to modulate their therapeutic efficacies in the context of atherosclerosis and cancer. This platform greatly facilitates the integration of existing organs-on-chip models to provide an intuitive and flexible way for users to configure different multi-organ perfusion systems.

The advent of 3D printing technologies promises to make microfluidic organ-on-chip technologies more accessible for the biological research community. To date, hydrogel-encapsulated cells have been successfully incorporated into 3D printed microfluidic devices. However, there is currently no 3D printed microfluidic device that can support multicellular spheroid culture, which facilitates extensive cell-cell contacts important for recapitulating many multicellular functional biological structures. Here, we report a first instance of fabricating a 3D printed microfluidic cell culture device capable of directly immobilizing and maintaining the viability and functionality of 3D multicellular spheroids. We evaluated the feasibility of two common 3D printing technologies i.e. stereolithography (SLA) and PolyJet printing, and found that SLA could prototype a device comprising of cell immobilizing micro-structures that were housed within a microfluidic network with higher fidelity. We have also implemented a pump-free perfusion system, relying on gravity-driven flow to perform medium perfusion in order to reduce the complexity and footprint of the device setup, thereby improving its adaptability into a standard biological laboratory. Finally, we demonstrated the biological performance of the 3D printed device by performing pump-free perfusion cultures of patient-derived parental and metastatic oral squamous cell carcinoma tumor and liver cell (HepG2) spheroids with good cell viability and functionality. This paper presents a proof-of-concept in simplifying and integrating the prototyping and operation of a microfluidic spheroid culture device, which will facilitate its applications in various drug efficacy, metabolism and toxicity studies.

The practical application of microfluidic liver models for in vitro drug testing is partly hampered by their reliance on human primary hepatocytes, which are limited in number and have batch-to-batch variation. Human stem cell-derived hepatocytes offer an attractive alternative cell source, although their 3D differentiation and maturation in a microfluidic platform have not yet been demonstrated. We develop a pump-free microfluidic 3D perfusion platform to achieve long-term and efficient differentiation of human liver progenitor cells into hepatocyte-like cells (HLCs). The device contains a micropillar array to immobilize cells three-dimensionally in a central cell culture compartment flanked by two side perfusion channels. Constant pump-free medium perfusion is accomplished by controlling the differential heights of horizontally orientated inlet and outlet media reservoirs. Computational fluid dynamic simulation is used to estimate the hydrostatic pressure heads required to achieve different perfusion flow rates, which are experimentally validated by micro-particle image velocimetry, as well as viability and functional assessments in a primary rat hepatocyte model. We perform on-chip differentiation of HepaRG, a human bipotent progenitor cell, and discover that 3D microperfusion greatly enhances the hepatocyte differentiation efficiency over static 2D and 3D cultures. However, HepaRG progenitor cells are highly sensitive to the time-point at which microperfusion is applied. Isolated HepaRG cells that are primed as static 3D spheroids before being subjected to microperfusion yield a significantly higher proportion of HLCs (92%) than direct microperfusion of isolated HepaRG cells (62%). This platform potentially offers a simple and efficient means to develop highly functional microfluidic liver models incorporating human stem cell-derived HLCs.

We present a first instance of performing 3D differentiation of a human liver stem cell in a pumpfree microfluidic platform. Pump-free perfusion culture at constant flow rate was achieved by using 3D printed supports to control the hydrostatic head difference between a pair of horizontally orientated media reservoirs connected to a microfluidic 3D cell culture device. We demonstrate that on-chip differentiation of HepaRG cells, a human bipotent liver stem cell, could efficiently generate functional hepatocytes and biliary cells, for up to 19 days.