A new approach to neural cell culture for long-term studies
Introduction
Dissociated primary neuron cultures have been a popular research tool for decades (Dichter, 1978, Mains and Patterson, 1973, Murray, 1965) because they allow easy access to individual neurons for electrophysiological recording and stimulation, pharmacological manipulations, and high-resolution microscopic analysis. It would be informative to follow individual cultured neurons for months, electrophysiologically and morphologically, to help understand how neural activity and morphology interact. In vivo or in vitro, it is technically difficult to record from and stimulate more than three cells using standard intracellular microelectrodes, and those cells usually die within minutes or hours. Given that neural systems use distributed codes to process and store information (Nicolelis et al., 1993, Wu et al., 1994), much of their dynamics is missed without a multi-unit approach. Multi-electrode arrays (MEAs) provide a means to record from and stimulate many individual cultured neurons non-destructively (Gross et al., 1982, Pine, 1980, Regehr et al., 1989). Because the array substrate consists of transparent glass, neuron morphology can be easily monitored using an inverted phase-contrast microscope, or using vital labels and a fluorescence microscope such as a confocal or 2-photon laser-scanning microscope (Potter et al., 1996, Potter et al., 2001).
A prerequisite to our program of studying long-term plasticity in cultured networks was to devise a culturing method that keeps neurons alive for many months. Why do neuron cultures die? One obvious cause of neuron death is infection, usually by mold. A less obvious, but equally prevalent one, is hyperosmolality due to medium evaporation (Maher and McKinney, 1995). Our method addresses both of these problems, and has allowed us to repeatedly study primary neuron cultures for >9 months, in one case for well over 1 year (Fig. 1). It involves growing cells on multielectrode arrays that are sealed by a selectively permeable membrane, in a non-humidified incubator.
Section snippets
Membrane-sealed multi-electrode culture chambers
Gas-tight culture chambers have previously been used in order to provide a more controlled environment for cultured cells (Kleitman et al., 1998, Morrison et al., 2000). It is necessary to fill such chambers with the proper mix of gases (usually air with 5% CO2) before sealing. Our approach is to let the incubator control the desired gas mixture, as it does in most labs, and use chambers that are permeable to the gases necessary for cell metabolism, but impermeable to microbes and water vapor.
Discussion
We have developed a method that enables the survival of primary neuron cultures for over a year in vitro. By sealing culture chambers with a membrane that is permeable to CO2 and O2, and relatively impermeable to water vapor, and by keeping these chambers in a non-humidified incubator, we have greatly reduced or eliminated problems with infection and hyperosmolality, while maintaining pH and O2 homeostasis. By combining this technique with multielectrode array dishes, it is now possible to
Sealed dish fabrication
Rings were machined from solid polytetrafluoroethylene (PTFE) Teflon® round stock to fit the MEAs tightly when a rubber O-ring (EP75, Real Seal, Escondido, CA) is fitted in the inside groove (see cross-section diagram, Fig. 2). A grove on the outside of the PTFE ring accommodates a second O-ring that holds on the membrane of fluorinated ethylene–propylene (Teflon® FEP film, 12.7 μm thickness, specified permeabilities to CO2, O2 and water vapor of 212, 95, and 78 micromol/cm2/day, respectively,
Acknowledgements
We thank Scott Fraser, Jerry Pine, Henry Lester, Sheri McKinney, Michael Maher, Sami Barghshoon, Alice Schmid, Axel Blau, and Daniel Wagenaar for their help and suggestions. This work was funded by the National Institute for Neurological Disorders and Stroke, grant RO1 NS38628.
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