As an organism with a short gestational period, the embryos of Drosophila melanogaster are prime study subjects for investigating the distribution and localization of proteins and their regulators during development. However, because of their lipid-rich embryos and chitin-rich chorion, their utility is limited by the difficulty of mounting embryos on glass surfaces. In this work, we introduce a practical method that significantly enhances the attachment of Drosophila embryo onto slides and detail our methods for successful histochemistry, immunohistochemistry, and in-situ hybridization.
Prior to the steps shown here, slides should be washed with detergent, then rinsed in tap water and distilled water before being placed in the oven to dry. Dry slides should gently soak in potassium dichromate for 24 hours before being rinsed and dried again, then placed in 95%ethanol for at least two hours. During this step, you can prepare further slides or the chrome alum-gelatin according to our methods.
Gelatin can then be stored in 60 degree bath water prior to use. To coat slides, drop a small amount of gelatin onto one edge of the slide. Then use another glass slide to slowly and evenly drag the gelatin across the surface of the slide to be coated.
This creates a thin, even layer. Before being put in a rack or drying apparatus to be stored in an oven to allow for full drying for another overnight or 48 hours. Embryo are collected using a grape juice agar plate, once ready, the grape juice agar plate can then be removed.
Flood the plate with PBS and gently brush with a soft brush in order to remove any attached embryos. Remove embryo containing PBS to a 1.5 milliliters centrifuge tube and allow to rest on ice. Once embryo are settled on ice, gently remove the supernatant PBS.
Be careful not to disturb the embryo at the bottom of the tube. Add 1 mL of 50%bleach to the sample and gently shake for two minutes. Remove the bleach by filtration and wash embryos in PBS three times.
Prepare n-heptane. Once ready, wash the embryos from the filter paper into n-heptane by shaking the filter paper gently in the solution. Allow embryos to settle at the bottom of the n-heptane plate and collect this n-heptane into a new 1.5 milliliter centrifuge tube.
Add one milliliter of methanol to the tube, allow the solution to precipitate and then aspirate the n-heptane methanol. Wash the embryo in PBS three to five times to remove residual methanol. Remove this extra PBS and add Bouin's fixative and allow to settle for 30 to 60 minutes in order to fix the embryos.
Remove Bouin's fixative and wash the embryos again in PBS three times. Prepare 1.2%agarose in PBS and keep warm in a 60 degree water bath then prepare the embryos for embedding by gently aspirating extra PBS that it has been stored in. Careful not to disturb the embryos at the bottom, and then aspirate the embryo plug at the bottom of the solution and place gently into a bottle cap mold.
Aspirate any additional PBS from this mold before adding 1.2%agarose gel. Prepare a solution of PBS in order to store these agarose plugs. Once the agarose has solidified, individual plugs can then be moved into this PBS for storage.
Here we are showing the sequential dehydration process. Allowing 15 minutes for incubation each time. Then remove from the 100%ethanol and fix in a one-to-one solution of ethanol and xylene for another 15 minutes.
Once removed from this one-to-one solution, remove in pure xylene for one minute. Now the tissue sample is ready to be moved into a warmed one-to-one xylene paraffin solution. Allow to sit for 30 minutes before final embedding in 100%paraffin.
Move to the first of 100%paraffin and rest for two hours. Repeat this process in its two and three paraffins. The sample is now ready to be moved into a mold and the mold filled with wax to complete the embedding process.
Embedding of Drosophila embryos. Place prepared paraffin sections of Drosophila embryos in a 60 degree oven for 15 minutes. Place them in a 100%solution of xylene.
We then begin rehydration. Place the slides in each of these solutions for three minutes each. Once the slide is in distilled water, place in the hematoxylin solution for two minutes.
Once complete make sure to drain as much of the solution as possible from the slide. Once the slide has been washed in distilled water, then stain the slide in eosin for one minute. Finally, remove the slide from the eosin and wash in distilled water.
Once this slide is clean, we now begin our final dehydration process. Finally being placed into 100%xylene for five minutes. This process is repeated in two new solutions of 100%xylene and drop a small amount of neutral gum along one edge of the slide.
Take this cover slip and gently position on one edge of the slide. First deparaffinize the slides by placing in 100%xylene. Once the slides are removed from distilled water, they should be covered with 1%periodic acid solution.
We then discard the excess solution and rinse in distilled water. By placing the slide directly into a preheated solution. Once this is complete we rinse the slide in distilled water and then tint the solution with 0.2%gold chloride for one to two minutes.
Next, we fix the tissue with 3%sodium thio sulfate for three minutes using distilled water. And then counterstain with hematoxylin. After counterstaining, the stain can be washed with either tube of distilled water as shown here.
Then we move on to the dehydration process. Follow this by putting the slides directly into 100%xylene and then apply a small amount of neutral gum to the corner of the slide. Gently cover with a cover slip.
Begin preparing the slides as previously by deparaffinizing and xylene. And then in one-to-one xylene ethanol. Once the slides are deparaffinized, we begin rehydrating their tissue and then finally in PBS.
Place the slides in 0.3%hydrogen peroxide made in methanol for 10 minutes. Then the slides are covered with 20 mLs of target retrieval solution. The slides can then be placed in the staining box.
Place the stainer box into a steamer, ensuring that no solution escapes and the slides are still entirely covered. Note that the stain box lid should not be entirely closed so that water vapor can escape during the steaming process. And then gently rinsing in PBS-T three times.
Discard the excess solution and repeat the process, this time with PBS. Next, we incubate the slides in a peroxide block for five to 10 minutes and then rinse several times again in PBS-T and then in PBS. Prepare the antibody and drop directly onto the tissue.
These tissues can then be incubated or at room temperature for four hours. The secondary antibody should be incubated at room temperature for 90 minutes, which is staining with 5%DAB for two to 10 minutes. After the appropriate color is obtained, rinse in distilled water, and then counterstain in hematoxylin.
After this hematoxylin stain, the slides can then be sequentially dehydrated. Place the slides into 100%xylene and allow to soak for three minutes. Then drop small amounts of neutral gum onto one edge of the slide.
Once a gum has been placed, take the slide cover and place gently from one edge of the slide. Add one microgram of purified linearized template DNA to a sterile RNase-free reaction vial. Then add enough sterile RNase-free DEPC treated double distilled water to make a total sample volume of 13 microliters.
Then centrifuge briefly and incubate for two hours at 37 degrees Celsius. For RNase protection experiments, we recommend incubating for 15 minutes with two microliters of Dnase-1, RNase-free to remove template DNA. Once incubation step is complete, Stop the reaction by adding two microliters of 0.2 molar EDTA at pH of eight.
Prior to the steps show here the slides were deparaffinized and rehydrated, according to our previously demonstrated protocol. After rehydration incubate the slides in 0.01 mole per liter PBS at pH of 7.4 for five minutes each to remove the fixative solution from the tissue. After rinsing in PBS, we next rinse with 100 millimolar per liter glycine solution for five minutes twice to remove free aldehyde groups from the tissue.
After this step, rinse slides with PBS for five minutes. Next, incubate in PBS containing 0.3%triton X-100 for 15 minutes prior to rinsing and then incubating again with one microgram per microliter of proteinase K.Next, fix slides with 4%paraformaldehyde and PBS at pH of 7.2 for three minutes. Then, wash the slides before incubating with 0.1 molar triethanolamine buffer pH 8, containing 0.25%acetic anhydride.
You may choose to tilt the box at this stage to ensure full coverage with the solution. The box is prepared by adding 20%glycerol or DEPC water to the bottom of a dry hybridization box. Next, cover the slides in 20 microliters of pre hybridization solution.
Salmon's sperm DNA at a concentration of 100 micrograms per milliliter to each slide. And then incubate at 42 degrees Celsius for four hours. This is done in a preprepared humidified box to ensure that the solution does not dry during this process.
In the post hybridization stage, it's very important that the hybridization solution is completely rinsed from your samples. This can be done in consecutive rinsings of 15 minutes at 37 degrees, using four times concentration SSC, two times concentration one times concentration, 0.5 times concentration and 0.1 times concentration SSC solution. Each incubating for 15 minutes at 37 degrees Celsius.
After this is done, wash with PBS three times for five minutes each before adding the washing buffer. Washing buffer should be rinsed for one minute prior to the next steps Ensure the washing buffer is completely dry from the slides before adding the next solution. Add 100 milliliters of blocking solution to each slide and incubate the slide for 30 minutes.
This is best done in a light free environment to ensure good staining. Next incubate the slides for 30 minutes again in 20 mLs of antibody solution. Apply the antibody solution directly onto slides for full coverage of your tissue.
Once incubation is complete, apply 100 milliliters of washing buffer. Incubate for 15 minutes in light free conditions to remove unbound antibody conjugates. Remove the washing buffer.
Next, apply 20 milliliters of detection buffer and allowed to incubate for approximately four minutes. After four minutes, remove the detection buffer. Dry the slides to ensure no additional solution and add 10 milliliters of freshly prepared color substrate solution.
This reaction begins within a few minutes, but it's complete after 16 hours. So allow to incubate for that period of time in a light free condition. Once the desired spots where bands are detected wash the slide with 50 milliliters of TE buffer to stop the reaction.
And then complete our previously shown dehydration process and mounting process for neutral gum. Figures one and two demonstrate the chrome aluminum gelatin coating method for paraffin sections and the embedding method for expression profile of FKBP12 and DmFKBP12 in developing Drosophila embryos. Figure three shows the protein distribution of antibody detected FKBP12 at the syncytial cellular blastoderm stages.
From these histochemical results, it can be seen that FKBP12 expression is less polarized in the syncytial blastoderm, then begins to localize within the basement membrane underneath the trophectoderm epithelium informs the gradient with more expression found in the anterior and posterior poles. Figure four, which follows the early and late gas relation stages of the Drosophila embryo shows FKBP12 protein becoming restricted to certain architectural components of the primitive embryonic tissue. With less extracellular distribution than that observed in early and late gastro embryonic stages.
Interestingly, the above described dynamic protein distribution is not accompanied by corresponding changes in MRI levels of FKBP12 as seen in figure five. Indicating protein expression to be the result of a still unknown post transcriptional molecular mechanism. Ryanodine receptor mediated calcium signaling is a fundamental pathway in many physiologic and pathologic processes of both vertebrate and invertebrate animals.
Mutations in this gene are known to cause many physiologic disturbances leading to disease or early cardiac death. However, the role this gene plays in early Drosophila development has been difficult to study. As a transit component of the embryo, lipid-rich embryo and chitin-rich chorion, make studies of protein and embryonic expression in early Drosophila development difficult.
The slide coating, embryo embedding, and immunohistochemical techniques described in this paper have allowed us to study in detail the dynamic distribution of ryanodine receptor protein and mRNA. We share these techniques and the hope that they will allow others to study other proteins during early development stages of Drosophila.
