This method can help answer key questions regarding the genetic regulation of the formation and maintenance of the blood-brain barrier during multiple stages of drosophila development. This method can also be adapted to examine the permeability of other barriers, like the intestinal epithelium in a variety of organisms. The main advantage of this technique is that it allows evaluation of the blood-brain barrier function in live tissues.
Visual demonstration of this method is critical, as the sample manipulations can be difficult and many steps require close attention to detail. For embryo collection place a minimum of 50 virgin females with 20 to 25 males per bottle with Corn Meal Agar food and incubate these flies for one to two days before beginning the collections. The day before the collection, warm apple juice agar plates at 25 degrees Celsius overnight.
The next morning transfer the carbon dioxide anesthetized flies to a collection cage and place a pre-warmed apple juice agar plate with a small smear of yeast paste on the open end of the cage. Then secure the plate to the cage with the red sleeve. To clear older embryos, allow the flies to lay eggs on an apple juice agar plate for one hour at 25 degrees Celsius.
At the end of the incubation, invert the cage mesh side down and tap the flies to the bottom of the cage. Replace the apple juice agar with a new pre-warmed apple juice agar plate with a small smear of yeast paste. Allow the flies to lay eggs on the new plate for another hour at 25 degrees Celsius.
To collect late stage 17 embryos for injection, replace the apple juice agar plate with a new pre-warmed plate with a smear of yeast paste. And allow the flies to lay eggs on the plate at 25 degrees Celsius. After one hour, replace the plate and age the plate with collected embryos for 19 hours at 25 degrees Celsius so that the embryos will be 20 to 21 hours old at the time of imaging.
At the end of the 19 hour incubation, remove the embryo collection plate from the incubator and cover the surface of the plate with PbTx. Use a paintbrush to loosen the embryos from the plate surface into a 70 micrometer nylon mesh strainer. And place the strainer with embryos in 50%bleach solution in a 100 millimeter Petri dish for five minutes with occasional agitation at room temperature to dechorionate them.
At the end of the incubation, rinse the embryos three times by swirling the strainer in fresh PbTx, using a new Petri dish for each wash. Wash embryos to one side of the strainer with PbTx and use a glass pipette to transfer the embryos onto a 2%agarose gel slab. Remove the excess liquid with filter paper.
Align six to eight embryos on the slab with the posteriors to the right and the dorsal sides facing up and firmly press a piece of double-sided tape affixed to a slide on top of the embryos. Then desiccate the embryos at room temperature for about 25 minutes before covering the specimens with Halocarbon oil. For larval collection, set up a cross with 5 to 10 virgin female flies of the desired genotype and half as many males of the desired genotype in a vial with Cornmeal Agar food.
After five to seven days at 25 degrees Celsius, depending on the genotype, use forceps to gently collect wandering third instar larvae from the vial and rinse the larvae with PBS to remove any stuck food. Transfer the larvae to an apple juice agar plate for genotyping as necessary before using a paintbrush to dry the larvae on a tissue. Then use forceps to transfer six to eight larvae to a slide prepared with double-sided tape.
Before the injection, use a micropipette puller to pull capillary tubes into a standard needle shape for D.Melanogaster injections and anchor the needles in clay in a Petri dish. For embryo injection, use a 20 microliter gel-loading pipette tip to load a prepared needle with 5 microliters of 10 kilodalton dextran conjugated to sulforhodamine 101 acid chloride and load the needle into a needle holder. Position the needle in a micromanipulator secured to a steel base and set the inject apparatus to 50 pounds per square inch and 5 to 10 milliseconds with a range of 10.
Place the slide on the micromanipulator stage and brush the edge of the needle against the edge of the double-sided tape at a 45 degree angle to create a slightly angled tip broken just enough to allow flow of the 10 kilodalton dextran through the opening. Pump the foot pedal until the dye is at the tip of the needle. Align the needle so that it is parallel with the embryo and puncture the posterior end of the specimen.
Pump the foot pedal to inject two nanoliters of dye into the specimen. Note the time of injection for incubation purposes and continue down the slide to inject additional specimens. For larval injection, make a slightly larger angled opening than that which is used for injecting embryos and angle the needle slightly downward toward the specimen before injecting the larvae with 220 nanoliters similar to as demonstrated for embryonic specimens.
At the end of the 10 minute injection incubation, use a cotton tipped applicator to apply petroleum jelly on the right and left sides of the samples on the slide as a spacer. Position the cover slip on top and place the slide onto confocal microscope stage. Select the 20X objective and image the samples throughout the depth of each embryo.
Then calculate the percentage of samples with a compromised blood-brain barrier according to the formula. Before beginning the dissection procedure, use nail polish to mount two cover slips spaced approximately 0.5 centimeters apart on a slide and place an injected larva into PBS on the slide at the end of the 30 minute incubation. Using one pair of forceps, grasp the larva halfway down the larval body and use a second pair of forceps to separate the anterior and posterior halves of the larva.
Next, use one pair of forceps to grip the anterior region at the mouth hooks and use the second pair of forceps to invert the body wall over the tip of the first pair of forceps. The brain and ventral nerve cord will be exposed. Separate the brain and ventral nerve cord from the body wall by severing nerves if needed and remove the body wall from the slide.
Cover the sample with 10 microliters of 80%glycerol. Then place a cover slip on top of the sample for imaging and image the samples to determine the percentage of samples with a compromised blood-brain barrier as demonstrated. When wild type late stage 17 embryos are injected with 10 kilodalton dextran conjugated to sulforhodamine 101 acid chloride fluorescent dye, the large dextran molecule is excluded from the ventral nerve cord, as expected.
Heterozygous raw 1 mutant embryos exhibit an intact blood-brain barrier, similar to that observed in wild type control embryos. In contrast, homozygous raw 1 mutant embryos exhibit defects in the integrity of the blood-brain barrier, with the 10 kilodalton dextran flooding into the ventral nerve cord, indicating a failure of the blood-brain barrier to form. In wild type control larval samples, 10 kilodalton dextran fails to penetrate the blood-brain barrier and is excluded from the brain and ventral nerve cord.
When attempting this procedure, proper aging of the embryos is critical to ensure the samples are examined at an age at which blood-brain barrier formation is expected to be complete. Following this procedure, mutants exhibiting a compromised blood-brain barrier can be furthered studied using genetics, immunohistochemistry, and microscopy to better understand gene function at the cellular level. This technique will allow researchers to identify additional genes that are critical for the establishment and maintenance of the blood-brain barrier.
