The overall goal of this Drosophila injury protocol is to observe both the preservation of axonal and synaptic morphology, as well as synaptic function of axons and their synapses. These assays can also be used to characterize neuronal maintenance factors, study axonal transport, or analyze axonal organelles in intact axons. The partial wing injury assay allows for the observation of injured axons undergoing degeneration side by side with uninjured control axons within the same nerve bundle in the Drosophila wing.
This injury assay can readily be practiced beforehand in wild-type flies by applying a cut roughly in the middle of the wing with micro-scissors. The internal injury allows for the assessment of axonal morphology and by the use of optogenetics to visualize functional preservation of axon and their synapses. Antennal ablation requires steady hands.
It is also recommended to practice the removal the antennae in wild-type flies beforehand. In addiction, any available optogenetics setup is suitable for activate the neurons in fly with hat. Otherwise, a simple setup can readily be built from scratch.
To begin, use five virgin females and five males from the right genotype to perform crosses at room temperature. Transfer the P0 generation into new vials every three to four days. Collect freshly eclosed adult progeny F1 generation daily into new vials and age them for seven to 14 days.
After anesthetization, use micro-scissors to cut the interior wing vein roughly in the middle of the wing. Use one wing for the injury and the other wing as an age matched uninjured control. Apply one injury per wing and make sure to get 15 wings injured.
Recover the flies in food containing vials. Next, with a pipette, spread 10 microliters of halocarbon oil 27 along a whole glass slide. One or seven days post injury, use micro-scissors to cut off the injured as well as the uninjured control wing.
Use tweezers to grab the wing at the center, mount maximal four wings into the halocarbon oil 27 and cover them with a cover slide. Images of GFP labeled neurons in the wing can readily be acquired with a spinning disk. However, the acquisition time has to be performed within less than eight minutes after mounting the wings because the dish is not fixed.
Image the wing immediately using a spinning disk microscope. Acquire a series of optical sections along the Z axis with 0.33 micrometer step size and compress Z-stacks into a single file for subsequent analyses. Cross five virgin females and five males from the right genotype and collect F1 generation as previously.
After anesthetization, use tweezers to oblate the right third antennal segment for unilateral ablation or both left and right third antennal segments for bilateral ablation. This removes GFP labeled neuronal cell bodies while their axonal projections remain in the CNS. Depending on the GFP labeled neurons used in the technique, it's important to know whether the cell bodies are housed in the third or in the second antennal segment for the subsequent injury assay.
Recover the flies in food containing vials. After anesthetization, use tweezers to grab the neck and another tweezers to fix the thorax. Gently pull the neck and head off the thorax.
Using tweezers that have been dipped into the fixing solution, transfer all heads into a 1.5 milliliter microcentrifuge tube containing one milliliter of fixing solution of 4%paraformaldehyde and 0.1 triton X100 in PBS. Fix the heads for 20 minutes with gentle agitation at room temperature. Then put the microcentrifuge tube on ice and the heads gravitate to the bottom of the microcentrifuge tube.
Remove the supernatant with a pipette and add one milliliter of washing buffer containing 0.1%triton X100 in PBS. Place the tube on a turning wheel at room temperature to wash for two minutes. Repeat the wash four additional times to remove residual fixing solution.
After preparing the brains, obtain a cover slide, stick lab tape on it, and cut out a T like shape from the tape. Use a 20 to 200 microliter pipette tip where three millimeters of the tip has been cut off to widen the opening of the pipette. Pipette the brain containing antifade reagent onto the slide and cover the brains with a cover slide.
Use clay to prepare two small even rolls. Ensure that the clay rolls are not higher than the height of a glass slide. Stick the clay rolls onto the glass slide and place the brain containing cover slide sandwich onto the clay rolls.
Proceed according to the manuscript. To prepare the flies for optogenetics, first melt fly food in a microwave. After the food cools down, before solidification, add 20 millimolar all trans-Retinal in ethanol to a final concentration of 200 micromolar.
Mix well and pour the food immediately into empty vials. Cover the vials containing the solidified food with plugs or cotton balls. Wrap the vials with aluminum foil.
Then, store the food containing vials in a dark cold room. Use five virgin females and five males from the right genotype to perform crosses at room temperature and collect F1 generation as previously in aluminum covered vials containing 200 micromolar all trans-Retinal in fly food. Next, collect the flies by tapping them from food containing vials into an empty vial with no food.
Cool the vial down in ice containing water for approximately 30 seconds. Flies fall asleep. Now, rapidly put individual flies into small chambers covered a cover slide to perform optogenetics.
In a dark room, perform the protocol consisting of 30 seconds where the red light is absent, followed by 10 seconds of red light exposure at 10 Hertz. Repeat this procedure three times in total followed by an additional 30 second interval where the red light is absent. Collect individual flies from each chamber on carbon dioxide pads.
To subject them to antennal injury, ablate both the left and right second antennal segments. This removes the cell bodies of Johnston's organ neurons while the axonal projections remain in the CNS. Recover the flies in aluminum covered vials containing 200 millimolar all trans-Retinal.
At corresponding time points, for example seven days post antennal ablation, subject the flies to another grooming assay. In this protocol, three methods were presented to study the morphology and function of severed axons and their synapses. The first method allows for high resolution observation of individual axons in the peripheral nervous system.
The schematic crosses to generate wild-type and highwire clones in the wing is shown here. The control in injured GFP labeled axons are also presented with arrows indicating severed axons. The second approach to study axon death of GFP labeled sensory neuron axons in the brain is presented.
Schematic crosses to generate wild-type and highwire clones in the brain is shown here. Examples of control in injured GFP labeled axons are presented with arrows indicating severed axon bundles. The third method demonstrates the approach to visualize axonal and synaptic function after axotomy.
The schematic crosses to generate wild-type and dnmnat over-expressing Johnston's organ sensory neurons is shown here. Wild-type flies are flies containing Johnston's organ neurons with attenuated axon death. Both harbored a potent grooming behavior before injury.
However, seven days post injury, grooming failed to be elicited by optogenetics in wild-type flies, while the animals with attenuated axon death continued to groom. Here we use loss of function mutations or our expression of genes to observe the preservation of axonal morphology or synaptic function. Other modifying methods can be applied, such as knock down RNA interference or tissue specific CRISPR-Cas9 mediated knock outs.
This methods can be used in an injury independent context. They allow for the observation and characterization of neuronal maintenance factors during aging, axonal transport, and axonal organelles in intact axons.
