Medicine
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A Drosophila In Vivo Injury Model for Studying Neuroregeneration in the Peripheral and Central Nervous System
Chapters
Summary May 5th, 2018
Here, we present a protocol using the Drosophila sensory neuron - dendritic arborization (da) neuron injury model, which combines in vivo live imaging, two-photon laser axotomy/dendriotomy, and the powerful fly genetic toolbox, as a platform for screening potential promoters and inhibitors of neuroregeneration.
Transcript
The overall goal of this Drosophila larval sensory neuron injury model is to combine in vivo live imaging, two-photon laser axotomy/dendriotomy, and the powerful fly genetic toolbox into a platform for screening potential promoters and inhibitors of neuroregeneration. This method will help address key questions in a field of neurogeneration, such as by identifying new intrinsic and extrinsic regulator for neurogeneration, both in a peripheral and central nerve system. The main advantage of this technique is that novel candidates for neuroregeneration can be screened in an easy, fast and cheap way.
Though this system can provide insight into neuroregeneration, it can also be applied to other systems, such as neurodegenerative diseases and neuron-glia interactions. Generally, individuals new to this method will struggle because different experimental setups that utilize different types of neurons are used to model axon versus dendritic regeneration. For the larvae collection, prepare culture bottles as follows.
Use a blade to punch a 1.5-centimeter hole in one wall of a Drosophila culture bottle, and fill the hole with a cotton ball for ventilation. Then put a dab of yeast paste on a grape juice agar plate, and use the plate to plug the main opening of the bottle. In such a bottle, set up a cross of 10 virgin females and five males, and change the plate daily while culturing at 25 degrees Celsius.
Culture the collected plates with a wet tissue soaked in mild propionic acid to prevent contamination. From the cultured plates, harvest larvae at the required stage using forceps. Gently transfer the picked larvae to a new grape juice agar plate without yeast paste.
After they have cleaned themselves off by crawling around, they can be imaged. To begin each imaging session, turn on the imaging lasers and the microscope. For the injury, use a two-photon microscope.
In the imaging software, set the laser to view GFP at 930 nanometers with a maximum power of 1950 milliwatts. Select Line Scan Mode, and open the pinhole all the way. Then increase the laser intensity to about 20%of full power to make a PNS injury, or to 50%to 100%of full power to make a VNC injury.
Next, select 512 square pixel frame for scanning, and use the highest possible scanning speed. Use an average number of one, and a bit depth of eight bits. Then set the gain to about 750, and set the offset to zero.
Now save this preset experimental protocol as 2P GFP 930 Ablation, allowing for easy reuse in further experiments. For post-injury imaging, use a confocal microscope. First set up the argon laser at 488 nanometers.
Select the Acquisition tab and then select Z Stack. Under Laser, turn on the 488 nanometer argon laser. Next go to Channels, select the 488 nanometer laser, and increase the laser power to five to 10%For the pinhole, use the option for one to two area units.
Then adjust the gain to 650. Now in Acquisition Mode, select the 1024 square pixels as the frame scan. Use the maximum scan speed.
Use an average number of two and a bit depth of eight bits. Save this pre-experimental protocol as GFP Imaging. Begin with anesthetizing the larvae.
In a fume hood, place a 60-millimeter glass dish into a 15-centimeter plastic petri dish. Then fold a piece of tissue paper and place it in the glass dish. Place the grape agar plate on the tissue, after the diethyl ether is added.
Next, onto a glass slide, place one drop of Halocarbon 27 oil at the center, and place a spot of vacuum grease on each of the four corners. Then use forceps to transfer one larva onto the agar plate, and cover the glass dish to anesthetize the larva. As soon as the larva stops moving, carefully transfer it to the Halocarbon oil, with its head upright.
Then place a cover slip over the slide and press it down gently until it touches the larva. Next, use gentle force to slide the cover slip to roll the cells to be ablated to where the two-photon laser will most easily hit them. The location will vary, depending on what neurons are being targeted.
Now secure the assembly on the two-photon microscope stage, and focus on the cells of interest, using a 40X oil immersion objective. In the software, switch to the scanning mode and load the saved protocol. Make sure the pinhole is opened all the way.
Then in Live Mode, get a good image of the region of interest. Next, stop the live scan so that the Crop button will become available. Using the Crop function, adjust the scan window to focus the target area on just the prospective site of injury.
Then open a new imaging window. Now reduce the scan speed and increase the laser intensity. Then toggle the Continuous button to start and stop the scan.
Watch carefully. As soon as there is a drastic increase in florescence, end the scan. Next switch back to the original imaging window and select the Live Mode, and find the region that was just targeted by adjusting the focus.
A good indication of successful injury is the appearance of a small crater, ring-like structure, or localized debris right on the injury site. If the laser power was too high, a large damaged area will be visible, which can be lethal. Now carefully remove the cover slip, and transfer the injured larva onto a new plate with yeast paste.
Put the plate into a 60-millimeter dish, along with a propionic acid soaked tissue. Then return the plate to culturing temperature. For subsequent imaging of the larva, make use of the saved confocal setup, and collect Z stack images with a 25X objective.
Be sure to include the normalization point, so the regeneration can be quantified. Using the described protocol, regeneration of class three and four DA neurons was investigated. Typically, three or four neurons on the right side of abdominal segments A7 to A2 were injured.
Specifically, class three DDAF and class four V Prime ADA neurons were targeted. Larvae were imaged at 24, 48, and 72 hours post-injury. At 24 hours, the distal axons usually completed degeneration, and the axon stem was readily visible.
At 48 hours, it was possible to assess regeneration in the class four DA neurons. About 70%of these neurons regenerated beyond the injury site. Even after 72 hours, it was clear that class three DA neurons, however, failed to regrow.
This was assessed based on repeated observations of stalled growth cones. After watching this video, you should have a good understanding of how to set up the experiment, perform two-photon injury, and assess the results. Once mastered, this technique takes 15 minutes per larvae, if it's performed properly.
While attempting this procedure, it's important to remember to compare the experimental growth with the corresponding controls, side by side. Following this procedure, a method like immunostaining can be performed in order to answer more questions, like whether axon injury can cause protein translocation, resulting in a pathway alteration. Since its development, this technique paved the way for the researchers in the field of neuroscience to explore neuroregeneration in Drosophila larval sensory neurons.
Finally, don't forget that working with diethyl ether can be extremely hazardous, and precautions, such as performing larval anesthesia in a fume hood, should always be taken while performing this procedure.
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