Immobilization of Live Caenorhabditis elegans Individuals Using an Ultra-thin Polydimethylsiloxane Microfluidic Chip with Water Retention

One advantage of this protocol is that the worm sheet is easy to handle. The sheet has features that prevent drying, enabling us to conduct microbeam irradiation to living animals, perform several behavioral assays, and long-term observation. The microfluidic channels in this worm sheet are open, making it easier to collect animals.

These worm sheets can also be used repeatedly, making them more economical. When we were developing this technique, we wanted to irradiate only a part of the animals with a microbeam under a microscope. So we needed clear, very thin chips capable of holding the animals.

To begin, place a thin, transparent sheet onto the stage for use as a bottom cover film. Gently position a worm sheet on the bottom cover film with flat tweezers. Place at least three droplets of buffer solution on the surface of a 6 centimeter, non-treated Petri dish.

Then, pick several adult C.elegans with a platina picker from a culture plate. Transfer the animals to a droplet with the platina picker and allow the animals to swim in buffer to remove any bacteria. Wash the animals in two separate droplets and rinse the animals in another droplet.

Next, place a droplet of buffer solution onto the surface of the worm sheet. Remove the washed animals from the buffer droplet, and transfer them to a droplet on the worm sheet. Use flat tweezers to place a PS cover film over the worm sheet, and press gently over the channels from one end of the sheet to the other.

Or simply drop animals into droplet and gently seal with the cover film so droplet spreads between the cover and the worm sheets with the animals within the channel. Confirm the animals are alive by checking for movement with a microscope at 1-2x magnification. Record the animals’positions and specifically note the number of the channel in which each animal’s enclosed, and the position of each animal in the channel.

For collecting immobilizing animals, use flat tweezers to remove the cover film from the worm sheet, then drop 10-15 microliters of buffer solution onto one of the microfluidic channels enclosing the animals. Under the stereomicroscope, observe the animals as they start to spin. Then use the platina picker to pick up the swimming animals and transfer them to an assay plate.

After enclosing the animals as previously demonstrated, observe fluorescent spots on the animals with a flourescence microscope. Then image calcium ion wave propagation using video acquisition to observe dynamic activities. Place the used worm sheet onto a six centimeter petri-dish and place about 100 microliters of sterilized ultra-pure water onto the sheet.

Use gloved fingers to spread water on the surface of the sheet and wash off any contaminants. Next, use disposable wipes to remove the moisture from the worm sheet. Dispense five milliliters of 70%ethanol into the Petri-dish and use gloved fingers to spread the ethanol over the worm sheet.

Finally, after the sheet is dry, place the sheet on a sterile Petri-dish and cover. Using this protocol, the suitability of different cover films was investigated. There was no significant difference in motility between control animals and animals enclosed in microfluidic channels with PS film.

In contrast, motility was significantly reduced in animals enclosed under a cover glass. The motility of animals enclosed with a pet film was also significantly decreased. It is important to select cover films for the sealing of microfluidic channels carefully.

Using this protocol, we evaluated the suitability of different cover films through locomotion assays of the animals three hours after on-chip immobilization. Using this technique, we intended to develop an ultra-thin wettable chip only for microbeam irradiation. However, these worm sheets have already been used for imaging and behavioral assays beyond their original applications.