Using Microfluidics Chips for Live Imaging and Study of Injury Responses in Drosophila Larvae

The overall goal of the following experiment is to non-invasively immobilize drosophila larvae for live imaging using a modified microfluidic chamber. A third in star larvae is positioned in the chip with some oil to help seal up the larvae. Next, the chip is placed on top of the larvae such that its body fits within the chamber.

The chip is connected to a syringe so that a vacuum can be applied. Motility of the larvae becomes restricted, and structures in the ventral part of the animal are pushed close to the cover slip. Using a confocal microscope, structures are easily imaged, such as the details of sensory neuron axon terminals, and regeneration of segmental nerves after induced nerve damage.

Other methods to immobilize josepha for live imaging, huge chloroform ether, or ISO anesthetics. The main advantage of this technique is that it avoids the huge of such toxins, which interfere with physiology of the animal. And added benefit from the absence of toxins is that the larva GB safe and to work with The robust immobilization allows for imaging of rapid events such as faster axonal transport and changes in intracellular calcium.

A single larva can be immobilized and imaged in the chip multiple times. This allows for time lives. Imaging of processes that take place over the course of up to 72 hours Begin by obtaining a sample PDMS chip to try out this protocol.

Instructions on how to make A-P-D-M-S chip from an SU eight mold can be read in the text protocol using a 21 gauge dispensing needle. Poke a hole in the vacuum port of the PDMS chip for an inverted microscope. Remove a 23 gauge needle from its base, the lock hub using a few twists.

Then insert the needle tip into a small piece of polyethylene tubing so that the tubing covers at least a millimeter of the needle. Use a razor blade to cut off the excess tubing. This leaves a plastic ring that can make a seal.

Insert the 23 gauge needle tip into the hole of the vacuum port. For an upright microscope, poke a second hole on the side of the PDMS chip with a 21 gauge dispensing needle. This hole will provide access to the first hole from the side.

Then insert the 23 gauge needle tip and the tubing ring into the side hole. Place a piece of double-sided tape over the top of the PDMS chip to seal the top hole. Attach a 20 centimeter length of polyethylene tubing between the needle tip inserted at the vacuum port of the chip and one of the ports of a three-way valve.

Then attach a 20 milliliter syringe into one of the two remaining ports of the valve leaving the last port open. Clean the PDMS chip with transparent adhesive tape. Attach a piece of tape to the bottom of the chip.

Make sure the tape is touching the entire PDMS surface, and then peel off the tape. Repeat this cleaning method three times. Since the PDMS chip is reusable, it is very important to remove oil residue as it can affect the adhesion of PDM to glass Transfer early foraging.

Third instar larvae from the food to a Petri dish containing water. It must be between 3.5 and four millimeters long to fit the chip correctly. Bathe the larvae to remove the culture medium.

Next plays a small drop of halo carbon 700 oil at the center of a glass cover slip using forceps gently pick up a washed larvae. Briefly place it on a lightweight wipe to towel it off, and then move it to the oil for 10 seconds. The drop should be small enough such that the trachea of the larvae are not coated.

Then place the larvae on a clean glass cover slip briefly and move it to another cover slip. Thus removing a bit more oil. Pay attention to the larvas orientation to image the neural cord and segmental nerves.

The ventral side should be downward. The readily identified tracheal tubes should be upwards. Now gently place the PDMS chip on top of the larvae.

Align the larvae to the center of the micro chamber. With its posterior oriented towards the vacuum port, the larva should not touch the edges of the chamber. Once aligned, push the PDMS chip against the glass cover slip to achieve a good seal.

Then double check that the larvae is entirely enclosed in the micro chamber. Now switch the three-way valve to the syringe. Hold the PDMS assemblage firm and pull the syringe plunger to withdraw two to 2.5 millimeters of air until resistance is felt.

Thus creating a vacuum. Then turn the valve off. To maintain the vacuum carefully, double check the larvae.

Its body should be immobilized inside the micro chamber, and the trachea should be visible. The rest of the PDMS chip should be in contact with the cover slip Orientations like these are not correct. Now image the larvae.

If using an upright microscope, fix the top side of the chip to the stage with double-sided tape. When the imaging is completed, release the vacuum. Then detach the PDMS chip from the cover slip.

The larva should be immediately motile using forceps. Return the larvae to a grape juice agar plate for recovery. Place a single anesthetized larva onto a grape juice agar plate.

Under the stereo microscope, turn the animal ventral side up. To visualize the ventral nerve cord and the segmental nerves, make sure that the larva is completely immobile. Locate the nerve cord and salivary glands.

It is important not to damage these structures. Locate the end of the third abdominal segment. Injury here damages the most nerves and is the easiest to reproduce without killing the animal.

The injury can also be conducted in more posterior segments using a duo star number five, forceps. Pinch the segmental nerves tightly through the cuticle for five to 10 seconds. When done correctly, the cuticle remains intact and the body wall is not pierced.

Mastery will take practice after the injury placed the animal ventral side down on the great plate. It should be able to move its head and eat. If the injury was successful, then the posterior half of the larvae will be paralyzed.

The larva chip was used to image the kinesin mediated transport of synaptic vesicles within individual peripheral axons. The entero grade movement of these vesicles was measured at about 1.0 microns per second. Retrograde movement was clocked at 0.8 microns per second.

The immobilization technique was used for laser microsurgery using a pulsed UV dye laser. Calcium levels were observed in specific neurons thanks to a genetically encoded indicator. G camp 3.0 a calcium spike is detected after a sensory neuron.

Dendrite is transected. Permitting the animal to rest between imaging sessions allows cellular events to be viewed over time. After crushing the axons of emeric motor neurons, the proximal axon stump underwent new sprouting.

Meanwhile, the distal axons formed varicosities and became fragmented through the process of wallerian degeneration. Using the larva chip tracking photo converted fluorescent proteins in vivo is also possible. A DRA two alpha tubulin fusion protein expressed in class four sensory neurons was photo converted in the cell bodies.

Within two days, a significant amount of photo converted protein was relocated to the axon terminals of the sensory neurons about a millimeter away from the original location in the cell body. After watching this video, you should have a good understanding of how to utilize the larvae chip for live imaging of drosophila larvae and how to conduct a nerve crush injury. Once these techniques are mastered, larvae can be positioned on the microscope in just a few minutes.

The nerve crush is just as quick. Therefore, these procedures can be utilized for large scale experiments, such as genetic degrees. The chip can be utilized to image structures on the ventral side of the drosophila larvae.

These include but are not limited to muscles, neuromuscular junction synapses, sensory neurons, peripheral nerves, and the ventral nerve cord. This procedure may also be used to conduct live imaging of the larval heart, salivary glands, and trachea. The larva chip used in this video is sized for animal between 3.5 and four millimeter in length.

For imaging is smaller or larger, animals chip with a smaller or larger chamber can be easily made. See the supplementary design file and instruction for making PDMS chips. Instructions for making the PDMS chips are included in the written protocol.

We can send you a sample chip if you contact us.