Neuroscience
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Visualization of the Axonal Projection Pattern of Embryonic Motor Neurons in Drosophila
Chapters
Summary June 16th, 2017
This work details a standard immunohistochemistry method to visualize motor neuron projections of late stage-16 Drosophila melanogaster embryos. The filleted preparation of fixed embryos stained with FasII antibody provides a powerful tool to characterize the genes required for motor axon pathfinding and target recognition during neural development.
Transcript
The overall goal of this experiment is to analyze the motor neuron projection patterns in late stage embryos of Drosophila melanogaster. This method can help answer key questions in the field of neural development, such as motor axon guidance and neural subcu formation. The main advantage of this technique is that it provides precise visualization of motor axon in developing embryos, which allows for reliable analysis of axonal pathfinding and target the condition defect.
One day after setting-up egg collection plates replace the plates with new ones containing a dab of freshly-made yeast paste. After three hours on the fresh plates, transfer the flies to new food bottles. And incubate the egg plates overnight at 25 degrees Celsius.
In the morning, add 1.8 milliliters of PBT solution to the plates. And use a cotton swab to dislodge the embryos. Then, using a 1 milliliter pipette, transfer the embryos and solution to a micro centrifuge tube.
Once the embryos settle to the bottom, aspirate as much solution as possible. Then rinse the embryos twice using one milliliter of PBT per rinse. To decoreonate the embryos, replace the last rinse with one milliliter of 50%bleach and rock the tube for three minutes at room temperature.
To remove the bleach, rinse the embryos three times with PBT. Then, replace the last rinse with a half milliliter of 100%heptane and a half milliliter of 4%paraformaldehyde. Now, incubate the embryos for 15 minutes with gentle rocking.
Following the incubation, remove the underlying solution layer and add back 500 microliters of 100%methanol. Now, for 30 seconds, shake the tube vigorously to devitellinize the embryos. Then, remove the overlying solution, add back 500 microliters of 100%methanol and shake the tube for another 10 seconds.
After the shaking the tube, tap it to help the embryos settle and remove as much solution as possible. Next, briefly rinse the embryos with 100%methanol. Then, immediately rinse the embryos with PBT three times.
Now, resuspend the embryos in one milliliter of fresh PBT. And incubate the embryos at 37 degrees Celsius for 15 minutes to prepare them for LacZ staining. While the embryos incubate, prepare a one milliliter aliquot of X-gal staining solution.
Warm the aliquot in a 65 degree Celsius water bath until it gets cloudy. Then transfer it to a 37 degree Celsius heat block. After the embryos have been warmed for 15 minutes remove the PBT solution, and replace it with the one milliliter aliquot of staining solution.
Then add 20 microliters of X-gal substrate. For about two to four hours, incubate the tube at 37 degrees Celsius with gentle rocking until a blue precipitate forms. Then, remove the staining solution and rinse the embryos three times with room temperature PBT.
After the rinses, use a needle probe or forceps to hand-sort the embryos under a dissection microscope. Collect the stained embryos of interest into a half millimeter micro centrifuge tube using a one milliliter pipette. Next, wash the selected embryos with 0.4 milliliters of PBT.
Then, replace the PBT with 0.3 milliliters of blocking solution and allow the embryos to incubate with gentle agitation for 15 minutes. After 15 minutes, add 75 microliters of the primary antibody and continue the incubation overnight. The following morning, wash the embryos with PBT for about two hours, changing the wash solution approximately every 30 minutes.
After the wash, add 0.3 milliliters of blocking solution containing the secondary antibody and incubate the tube with gentle rocking at room temperature overnight. The next morning, wash the embryos with PBT for about three hours. Change the PBT at least five times during the wash period.
After the wash, resuspend the embryos in 0.3 milliliters of dab solution and add two microliters of 3%hydrogen peroxide. Then, incubate them in the dark with gentle rocking at room temperature until enough precipitate is produced. This generally takes 30 to 60 minutes.
Then, rinse the embryos with PBT four times and resuspend them in 0.2 milliliters of mounting solution. Now, allow the embryos to equilibrate in the dark at 4 degrees Celsius. To filet the embryos, stage them on a glass slide.
First move them into a drop of glycerol ventral side up. Next, under a dissection microscope, cut off the anterior part and 1/4 of the body length and remove the most posterior region, which is free of motor axons. Now, using a needle probe, move the preparation out of the glycerol, oriented dorsal side up and position it horizontally in the field of view.
The next step requires a fine tungsten needle that has been inserted into the hollow region of a needle probe, and then bent at the end. Using the tungsten needle, make a small cut at the posterior end of the embryo and continue to cut down the dorsal midline. Make sure the tip of the tungsten needle does not touch the surface of the glass slide during the dissection, because the needle will easily bend against the glass.
Then, with the probe return preparation back to the glycerol drop and position it dorsal side up. There, detach the gut from the body wall by unfolding each body wall in a ventral-lateral direction. The two body walls should come apart.
Now using the probe, carefully move the body wall flaps onto a glass slide. On the slide, use a tungsten needle to remove the internal organs by pushing them laterally. To stabilize the tungsten needle and keep it from being damaged, keep the hollow needle probed and hold the tungsten needle braced against the surface of the glass slide while manipulating the tungsten needle.
Now, mount the preparations in eight microliters of mounting solution on a new glass slide. Attach a cover slip and seal the edges down with regular nail polish. Then, capture images at high resolution using DIC optics.
Using the described method, stage 16 embryos were stained with Fas2 antibody and prepared for visualization of the motor neurons and abdominal hemi-segments A3 and A4.Normally, at least seven motor neurons extend and fasciculate their axons to form an ISNb nerve branch. Many surrounding nerve bundles were also identifiable in this preparation. When the intersegmental nerve bundle reaches the choice point at muscle 6 and 7, two axons selectively de-fasciculate and enervate muscles 6 and 7.
The same occurs at other choice points as the bundle extends dorsally. Then, at the last choice point, two axons enervate muscle 12. By contrast, in Sema-1a mutant embryos the same axons fail to defasciculate at their choice points.
This is not unexpected as the Sema-1 aegean product is known to help form connections between motor axons and target muscles during neural development. Thus, the described method can be used to analyze mutant phenotypes. After watching this video, you should have a good understanding of how to sort the desired mutant embryos, how to immunostain the axonal projection patterns of motor neurons, and how to dissect fixed embryos for flat preparation.
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