Microfluidic platforms for the study of neuronal injury in vitro.

Traumatic brain injury (TBI) affects 5.3 million people in the United States, and there are 12,500 new cases of spinal cord injury (SCI) every year. There is yet a significant need for in vitro models of TBI and SCI in order to understand the biological mechanisms underlying central nervous system (CNS) injury and to identify and test therapeutics to aid in recovery from neuronal injuries. While TBI or SCI studies have been aided with traditional in vivo and in vitro models, the innate limitations in specificity of injury, isolation of neuronal regions, and reproducibility of these models can decrease their usefulness in examining the neurobiology of injury. Microfluidic devices provide several advantages over traditional methods by allowing researchers to (1) examine the effect of injury on specific neural components, (2) fluidically isolate neuronal regions to examine specific effects on subcellular components, and (3) reproducibly create a variety of injuries to model TBI and SCI. These microfluidic devices are adaptable for modeling a wide range of injuries, and in this review, we will examine different methodologies and models recently utilized to examine neuronal injury. Specifically, we will examine vacuum-assisted axotomy, physical injury, chemical injury, and laser-based axotomy. Finally, we will discuss the benefits and downsides to each type of injury model and discuss how researchers can use these parameters to pick a particular microfluidic device to model CNS injury.

Collagen nanofibre anisotropy induces myotube differentiation and acetylcholine receptor clustering.

To create musculoskeletal tissue scaffolds for functional integration into host tissue, myotubes must be properly aligned with native tissue and spur the formation of neuromuscular junctions. However, our understanding of myoblast differentiation in response to structural alignment is incomplete. To examine how substrate anisotropy mediates myotube differentiation, we studied C2C12 myoblasts grown on aligned collagen substrates in the presence or absence of agrin. Myoblasts grown on microfluidically patterned collagen substrates demonstrated increased multinucleated myotubes and nicotinic acetylcholine receptor (AChR) clusters. However, agrin treatment did not synergistically increase differentiation of myoblasts seeded on these patterned collagen substrates. Myoblasts grown on aligned electrospun collagen nanofibres also demonstrated increased formation of multinucleated myotubes and AChR clusters, and agrin treatment did not increase differentiation of these cells. Using fluorescently labelled collagen nanofibres, we found that AChR clustered in cells grown on nanofibres with significantly higher anisotropy and that this clustering was eliminated with agrin treatment. Interestingly, anisotropy of substrate had no effect on the localization of AChRs along the myotube, suggesting that additional signalling pathways determine the specific location of AChRs along individual myotubes. Taken together, our results suggest a novel role for fibre anisotropy in myotube differentiation, specifically AChR clustering, and that anisotropy may guide differentiation by activating similar pathways to agrin. Our data suggest that agrin treatment is not necessary for differentiation and maturation of myoblasts into myotubes when myoblasts are grown on aligned collagen substrates.

Microfluidic device-assisted etching of p-HEMA for cell or protein patterning.

The construction of biomaterials with which to limit the growth of cells or to limit the adsorption of proteins is essential for understanding biological phenomena. Here, we describe a novel method to simply and easily create thin layers of poly (2-hydroxyethyl methacrylate) (p-HEMA) for protein and cellular patterning via etching with ethanol and microfluidic devices. First, a cell culture surface or glass coverslip is coated with p-HEMA. Next, a polydimethylsiloxane (PDMS) microfluidic is placed onto the p-HEMA surface, and ethanol is aspirated through the device. The PDMS device is removed, and the p-HEMA surface is ready for protein adsorption or cell plating. This method allows for the fabrication of 0.3 µm thin layers of p-HEMA, which can be etched to 10 µm wide channels. Furthermore, it creates regions of differential protein adhesion, as shown by Coomassie staining and fluorescent labeling, and cell adhesion, as demonstrated by C2C12 myoblast growth. This method is simple, versatile, and allows biologists and bioengineers to manipulate regions for cell culture adhesion and growth. © 2017 American Institute of Chemical Engineers Biotechnol. Prog., 34:243-248, 2018.

A Versatile Method of Patterning Proteins and Cells.

Substrate and cell patterning techniques are widely used in cell biology to study cell-to-cell and cell-to-substrate interactions. Conventional patterning techniques work well only with simple shapes, small areas and selected bio-materials. This article describes a method to distribute cell suspensions as well as substrate solutions into complex, long, closed (dead-end) polydimethylsiloxane (PDMS) microchannels using negative pressure. This method enables researchers to pattern multiple substrates including fibronectin, collagen, antibodies (Sal-1), poly-D-lysine (PDL), and laminin. Patterning of substrates allows one to indirectly pattern a variety of cells. We have tested C2C12 myoblasts, the PC12 neuronal cell line, embryonic rat cortical neurons, and amphibian retinal neurons. In addition, we demonstrate that this technique can directly pattern fibroblasts in microfluidic channels via brief application of a low vacuum on cell suspensions. The low vacuum does not significantly decrease cell viability as shown by cell viability assays. Modifications are discussed for application of the method to different cell and substrate types. This technique allows researchers to pattern cells and proteins in specific patterns without the need for exotic materials or equipment and can be done in any laboratory with a vacuum.

Vacuum-assisted fluid flow in microchannels to pattern substrates and cells.

Substrate and cell patterning are widely used techniques in cell biology to study cell-to-cell and cell-substrate interactions. Conventional patterning techniques work well only with simple shapes, small areas and selected bio-materials. This paper describes a method to distribute cell suspensions as well as substrate solutions into complex, long, closed (dead-end) polydimethylsiloxane (PDMS) microchannels using negative pressure. Our method builds upon a previous vacuum-assisted method used for micromolding (Jeon et al 1999 Adv. Mater 11 946) and successfully patterned collagen-I, fibronectin and Sal-1 substrates on glass and polystyrene surfaces, filling microchannels with lengths up to 120 mm and covering areas up to 13 × 10 mm(2). Vacuum-patterned substrates were subsequently used to culture mammalian PC12 and fibroblast cells and amphibian neurons. Cells were also patterned directly by injecting cell suspensions into microchannels using vacuum. Fibroblast and neuronal cells patterned using vacuum showed normal growth and minimal cell death indicating no adverse effects of vacuum on cells. Our method fills reversibly sealed PDMS microchannels. This enables the user to remove the PDMS microchannel cast and access the patterned biomaterial or cells for further experimental purposes. Overall, this is a straightforward technique that has broad applicability for cell biology.