The sophistication and success of recently reported microfabricated organs-on-chips and human organ constructs have made it possible to design scaled and interconnected organ systems that may significantly augment the current drug development pipeline and lead to advances in systems biology. Physiologically realistic live microHuman (µHu) and milliHuman (mHu) systems operating for weeks to months present exciting and important engineering challenges such as determining the appropriate size for each organ to ensure appropriate relative organ functional activity, achieving appropriate cell density, providing the requisite universal perfusion media, sensing the breadth of physiological responses, and maintaining stable control of the entire system, while maintaining fluid scaling that consists of ~5 mL for the mHu and ~5 µL for the µHu. We believe that successful mHu and µHu systems for drug development and systems biology will require low-volume microdevices that support chemical signaling, micro-fabricated pumps, valves and microformulators, automated optical microscopy, electrochemical sensors for rapid metabolic assessment, ion mobility-mass spectrometry for real-time molecular analysis, advanced bioinformatics, and machine learning algorithms for automated model inference and integrated electronic control. Toward this goal, we are building functional prototype components and are working toward top-down system integration.

Real-time monitoring of changes to cellular bioenergetics can provide new insights into mechanisms of action for disease and toxicity. This work describes the development of a multi-analyte screen-printed electrode for the detection of analytes central to cellular bioenergetics: glucose, lactate, oxygen, and pH. Platinum screen-printed electrodes were designed in-house and printed by Pine Research Instrumentation. Electrochemical plating techniques were used to form quasi-reference and pH electrodes. A Dimatix materials inkjet printer was used to deposit enzyme and polymer films to form sensors for glucose, lactate, and oxygen. These sensors were evaluated in bulk solution and microfluidic environments, and found to behave reproducibly and possess a lifetime of up to six weeks. Linear ranges and limits of detection for enzyme-based sensors were found to have an inverse relationship with enzyme loading, and iridium oxide pH sensors were found to have super-Nernstian responses. Preliminary measurements where the sensor was enclosed within a microfluidic channel with RAW 264.7 macrophages were performed to demonstrate the sensors’ capabilities for performing real-time microphysiometry measurements.

Prior exposure to sub toxic insults can induce a powerful endogenous neuroprotective program known as ischemic preconditioning. Current models typically rely on a single stress episode to induce neuroprotection whereas the clinical reality is that patients may experience multiple transient ischemic attacks (TIAs) prior to suffering a stroke. We sought to develop a neuron enriched preconditioning model using multiple oxygen glucose deprivation (OGD) episodes to assess the endogenous protective mechanisms neurons implement at the metabolic and cellular level for stress adaptations. We found that neurons exposed to a five minute period of glucose deprivation recovered oxygen utilization and lactate production using novel microphysiometry techniques. Using the non-toxic and energetically favorable five minute exposure, we developed a preconditioning paradigm where neurons are exposed to this brief OGD for three consecutive days. These cells experienced 45% greater survival following an otherwise lethal event and exhibited a longer lasting window of protection in comparison to our previous in vitro preconditioning model using a single stress. As in other models, preconditioned cells exhibited mild caspase activation, an increase in oxidized proteins and a requirement for reactive oxygen species for neuroprotection. Heat shock protein 70 was upregulated during preconditioning, yet the majority of this protein was released extracellularly. We believe coupling this neuron enriched multiday model with microphysiometry will allow us to assess neuronal specific real-time metabolic adaptations necessary for preconditioning.

Multianalyte microphysiometry is a powerful technique for studying cellular metabolic flux in real time. Monitoring several analytes concurrently in a number of individual chambers, however, requires specific instrumentation that is not available commercially in a single, compact, benchtop form at an affordable cost. We developed a multipotentiostat system capable of performing simultaneous amperometric and potentiometric measurements in up to eight individual chambers. The modular design and custom LabVIEW™ control software provide flexibility and allow for expansion and modification to suit different experimental conditions. Superior accuracy is achieved when operating the instrument in a standalone configuration; however, measurements performed in conjunction with a previously developed multianalyte microphysiometer have shown low levels of crosstalk as well. Calibrations and experiments with primary and immortalized cell cultures demonstrate the performance of the instrument and its capabilities.

Multianalyte microphysiometry, a real-time instrument for simultaneous measurement of metabolic analytes in a microfluidic environment, was used to explore the effects of cholera toxin (CTx). Upon exposure of CTx to PC-12 cells, anaerobic respiration was triggered, measured as increases in acid and lactate production and a decrease in the oxygen uptake. We believe the responses observed are due to a CTx-induced activation of adenylate cyclase, increasing cAMP production and resulting in a switch to anaerobic respiration. Inhibitors (H-89, brefeldin A) and stimulators (forskolin) of cAMP were employed to modulate the CTx-induced cAMP responses. The results of this study show the utility of multianalyte microphysiometry to quantitatively determine the dynamic metabolic effects of toxins and affected pathways.