We present protocols for the collection, preparation, and imaging of mature Drosophila oocytes. These methods allow the visualization of chromosome behavior and spindle assembly and function during meiosis.
The overall goal of this procedure is to prepare Drosophila oocytes for the cytological examination of spindle assembly and chromosome orientation during meiosis I.This method can help answer key questions in the meiosis field, such as how spindle bipolarity is established and how chromosomes become bi-oriented in the absence of centrosomes. The main advantage of this technique is that the oocyte membranes that block antibody penetration are effectively removed, which allows immunofluorescence. To begin pre-coat the inside of a five-millimeter tube and pasture pipette with PTB to prevent oocytes sticking to the surfaces.
Add 100 milliliters of ROB's buffer to a blender and then add 100 to 300 anesthetized flies and pulse three times for one second. Filter the resulted slurry into a 250-milliliter beaker through a large mesh with pores of around 1, 500 micrometers. Let the filtrate settle for around two minutes and then aspirate the top layer, removing as many large body parts as possible.
Filter the slurry again into a 250-milliliter beaker, through a smaller mesh with the pore size of 300 micrometers. Rinse the first beaker using additional buffer in the coated pipette to collect the remaining oocytes. Let the slurry settle for three minutes and then aspirate all but the bottom 10 milliliters.
Pour as much of the 10 milliliters as possible into the coated five-milliliter tube. Let the slurry settle for around two minutes and then carefully remove the liquid. Add the remainder of the 10 milliliters of slurry and let the mix settle again before removing the liquid.
Rinse any remaining oocytes out of the beaker using additional buffer in the coated pipette. Let the rinse settle in the five-milliliter tube for three to five minutes. To fix the ovaries, first aspirate off all liquid from the tissues and immediately add one milliliter of Fix Mix.
Place the tissues on a nutator for 2 1/2 minutes. Next, add one milliliter of heptane, vortex the tube for one minute, and then allow the oocytes to settle for a further minute. Remove all liquid from the settled tissue, then add one milliliter of 1x PBS.
Vortex the tube for 30 seconds and then allow the oocytes to settle for one minute. After that, remove all liquid from the tube and fill it with five milliliters of 1x PBS. Using the coated pipette, add roughly 500 to 1, 000 oocytes to a frosted glass slide.
Remove any extraneous body parts or material using forceps. Place a cover slip on top of the tissue and gently roll the oocytes until all membranes are removed. Dragging the edge of the coverslip across the oocytes works well.
Check the progress of membrane removal under a microscope periodically. Too much pressure will destroy the oocytes. The most effective oocyte membrane removal is achieved by combining the maximum amount of oocyte movement with the minimum amount of pressure.
Use a light touch and quick movements to drag the edge of the coverslip across the oocytes. To begin extraction and blocking, add 15 milliliters of PBS 1%Triton X-100 to a 50-milliliter tube. Rinse the rolled oocytes into this tube and nutate for 1 1/2 to two hours.
After allowing the oocytes to settle for two minutes, remove the liquid along with as many membranes as possible and add 50 milliliters of PBS 0.05%Triton X-100. Let the oocytes settle for a further two minutes, and then remove all but around one milliliter of liquid. Transfer the oocytes to a graduated 1.5 milliliter tube and remove the remaining liquid.
Add one milliliter of PTB for blocking and nutate for one hour. Remove the liquid from the oocytes and then add 300 microliters of PTB. Next, add the primary antibody of interest at the appropriate dilution, taking care to protect the samples treated with fluorescent antibody from light.
Nutute overnight at four degrees Celsius. Wash the oocytes and one milliliter of PTB four times for 15 minutes each. Remove the liquid, and add 200 microliters of supernatant from the secondary antibody-treated embryos.
Filter the 300-microliter mark with PTB and then nutate for three to four hours at room temperature. Wash the oocytes once for 15 minutes with one milliliter of PTB. Next, remove the liquid and add 0.5 microliters of Hoechst 33342 and 500 microliters of PTB.
Set to nutate for seven minutes. Use one milliliter of PTB to wash the oocytes two times for 15 minutes each. The samples can now be mounted and imaged immediately using standard confocal microscopy techniques, or stored in PTB at four degrees Celsius until needed.
Images of late-stage oocytes generated using the demonstrated technique are shown here, capturing three different stages of meiosis. Oocytes in prophase are distinguished by the presence of the nuclear envelope which is visible due to the lack of tubulin signal in the regions surrounding the karyosome. The distinctive elongation of the karyosome in the prometaphase was also observed, and the fourth chromosomes are seen apart from the karyosome mass.
Metaphase was also captured and the karyosome is seen retracted into a round shape. Here immunostaining with different antibodies highlights the spindle, centromeres, and central spindle in oocytes fixed with formaldehyde and heptane. Oocytes treated with the drug Colchicine showed non-standard phenotypes with the elimination of most non-kinetic or microtubules, resulting in all microtubules contacting the DNA at centromere labeled Phocyde.
Treatment with Paclitaxel resulted in oocytes with excessive microtubules in the cytoplasm. Finally oocytes treated with Binucleine 2 to inhibit Aurora B kinase, showed complete loss of spindle microtubules. While attempting this procedure, it's important to remember not to rush during oocyte membrane removal, because the success of the protocol depends on this step.
Generally individuals new to this method will struggle, because the removal of the membranes requires patience and practice. This technique is therefore best learned by watching someone perform it successfully. Once mastered, rolling the oocytes can be done in five to 10 minutes.
But it's possible to take up to 30 minutes without affecting the quality of the results. After watching this video, you should have a good understanding of how to prepare late-stage Drosophila oocytes for imaging biotic spindle assembly, spindle organization, chromosome movements, and chromosome bi-orientation. Thanks for watching, and good luck with your experiments.
