Using Fluorescence In Situ Hybridization (FISH) to Monitor the State of Arm Cohesion in Prometaphase and Metaphase I Drosophila Oocytes

The overall goal of this procedure is to generate FISH probes and visualize the state of meiotic arm cohesion in drosophila oocytes during prometaphase one and metaphase one. The main advantage of this technique is that it allows one to assay the state of arm cohesion in prometaphase and metaphase one arrested drosophila oocytes. This method can help answer key questions in the field of drosophila meiosis such as what conditions lead to premature loss of arm cohesion?

This technique has the potential to expand our understanding of the maternal age effect in humans which occurs, at least in part, because meiotic cohesion deteriorates as oocytes age. Begin this protocol with the generation of the arm probe for FISH, as well as the dissection and fixation of oocytes as detailed in the text protocol. To separate the late-stage oocytes, first add one milliliter of PBSBTX to a shallow dissecting dish.

Then, use a P-200 with a BSA-coated tip to transfer fixed ovaries into the shallow dish. Pipette the ovaries up and down with the BSA-coated pipette tip to dislodge the mature oocytes from the less mature oocytes. When late-stage oocytes are sufficiently separated, transfer all the tissue to a 500-milliliter microfuge tube.

Remove excess liquid with a pulled Pasteur pipette leaving about 150 to 200 microliters in the tube. To prepare for oocyte rolling, pre-wet a deep-well dish with 200 microliters of PBSBTX. Cover the dish and set it aside.

Obtain three frosted glass slides, and set slide three aside. Next, gently rub the frosted glass regions of slides one and two together. Rinse them in deionized water to remove any glass shards and dry with a disposable wipe.

Coat the frosted regions of slides one and two with PBSBTX by adding 50 microliters of PBSBTX to one slide and rubbing this region with the other slide. Remove the liquid a disposable wipe, and place the slides under a dissecting microscope. Keep the frosted regions of slides one and two in contact, with slide three supporting slide two.

To roll the oocytes, first pre-wet a P-200 pipette tip in PBSBTX and disperse the oocytes in the microfuge tube by pipetting up and down. Transfer 50 microliters of liquid containing the oocytes to the center of the frosted glass part of slide one. Lift slide two to do this.

Slowly lower slide two until the surface tension of the liquid creates a seal between the two frosted glass regions. There should be enough liquid to cover the frosted area, but none should be seeping out. Then, hold the bottom slide one in place with one hand, and use the other hand to move the top slide two back and forth in a horizontal direction keeping slide two level and supported on slide three.

Perform under a microscope for easy visualization of oocyte movements and progress. After a few movements in the horizontal direction, slightly change the angle of movement. In multiple increments, gradually increase this angle to 90 degrees until movement of the top slide two is perpendicular to the starting direction.

Note that empty chorions will be visible in the liquid and oocytes lacking chorions will appear longer and thinner. When rolling the oocytes, ensure that the direction of rolling is always in a straight line and never a circular motion. Repeat the rolling about seven to 10 times until the solution becomes slightly cloudy.

Stop rolling when the majority of oocytes appear to have lost their vitelline membranes. Gently lift the top slide two dragging one of its corners to the center of the frosted region of the bottom slide one so that the rolled oocytes accumulate in the center of the frosted region. Rinse the oocytes from both slides with PBSBTX into the deep-well dish containing PBSBTX.

Clean slides one and two with ultrapure water. Dry with a disposable wipe and reset. Repeat these steps until all the oocytes of the same genotype have been rolled.

This usually requires three to four rounds of rolling per genotype. To remove debris after rolling, add one milliliter of PBSBTX to a 15-milliliter conical tube. Swirl the liquid to coat the sides of the tube.

Using a PBSBTX-coated P-1, 000 pipette tip, transfer the rolled oocytes from the deep-well dish to the conical tube containing one milliliter of PBSBTX. Add an additional two milliliters of PBSBTX to the conical tube containing the oocytes. Hold the conical tube against a dark background to see the opaque oocytes as they sink.

After letting the oocytes settle to the bottom, use a P-1, 000 to remove the top two milliliters of solution containing debris and discard. After repeating the step for a total of three rounds of debris removal, use a PBSBTX-coated P-1, 000 pipette tip to transfer the oocytes back to the original 500-microliter microfuge tube. 20 to 25 females should yield approximately 50 microliters of rolled mature oocytes.

Repeat this process for the remaining genotypes using fresh frosted slides, a clean deep-well dish and a new conical tube for each genotype. Rinse the oocytes with 500 microliters of PBS containing 1%Triton X-100. Remove the liquid and add 500 microliters of extraction buffer.

Incubate the sample on the nutator at room temperature for two hours. After performing pre-hybridization washes as described in the text protocol, perform the denaturation and hybridization. First, use a BSA-coated P-200 pipette tip to transfer the oocytes to a 200-microliter PCR tube.

Keep samples at 37 degrees Celsius and let the oocytes settle to the bottom. It is important to be patient when changing solutions so that the oocytes are not lost in the process. Once the oocytes have settled, use a pulled Pasteur pipette to remove as much liquid as possible.

Then, add 40 microliters of probe-containing hybridization solution to the oocytes. Place the PCR tube containing the oocytes in the PCR machine and incubate as listed in the text protocol. After the probe is added to the oocytes, keep them in the dark as much as possible.

For post-hybridization washes, preheat the 2X SSCT plus 50%formamide to 37 degrees Celsius and keep it in the hybridization incubator. With a BSA-coated P-200 pipette tip, add 100 microliters of 37-degrees Celsius 2X SSCT plus 50%formamide to the PCR tube with the oocytes, and transfer the oocytes to a new 500-microliter microfuge tube. After adding an additional 400 microliters of 37-degrees Celsius 2X SSCT plus 50%formamide to the same tube, let the oocytes settle at 37 degrees Celsius in the incubator.

Settling often takes longer at this step. It is not unusual to take 15 minutes or more. A lack of patience will result in a significant loss of oocytes.

After allowing the oocytes to settle, remove liquid with a pulled Pasteur pipette. Add 500 microliters of 37-degrees Celsius 2X SSCT plus 50%formamide at 37 degrees Celsius and wash the oocytes with rotation. Following each wash slash rotation step, keep the oocytes at 37 degrees Celsius while they settle.

After additional washes at room temperature, stain with DAPI by washing oocytes for 20 minutes in 500 microliters of one-microgram-per-milliliter DAPI solution in 2X SSCT on the nutator. After rinsing the oocytes three times in 500 microliters of 2X SSCT, wash the oocytes two times for 10 minutes each in 500 microliters of 2X SSCT on the nutator. Samples can be stored for up to four hours at room temperature in the dark until they are ready to mount on the cover slips.

With a BSA-coated P-200 pipette tip, pipette up and down to disperse oocytes throughout the solution. Then, transfer 40 microliters of the oocyte solution to a poly-L-lysine-coated cover slip. Remove some of the liquid from the cover slip using a pulled Pasteur pipette until the oocytes stick to the cover slip but are still surrounded by liquid.

With forceps, hold down the cover slip on one side, and use a Tungsten needle to gently dissociate clumps of oocytes moving them to achieve a single layer of non-overlapping oocytes. Remove the remainder of the liquid around the oocytes with a pulled Pasteur pipette. Then, slowly lower the slide towards the cover slip with the mounting media facing the sample until the media touches the sample.

After letting the slides dry for three to five days in the dark, acquire laser scanning confocal images using a high-numerical aperture 40 times oil objective with six times digital zoom as detailed in the text protocol. A software package that allows one to view images and score for cohesion defects in three dimensions is imperative. This allows one to scroll through the Z series to view the FISH spots and to accurately count the number of individual arm signals.

Maximum intensity projections of confocal Z series are shown for oocytes with intact and disrupted arm cohesion. In these figures, the arm probe signal is yellow and the pericentric probe signal is red. Two arm probe spots indicate that sister chromatids are together and arm cohesion is intact.

Three arm probe spots indicate that one pair of sister chromatids have prematurely lost arm cohesion while the other pair of sisters has not. If two arm probe spots are connected by a thread, the sister chromatids are not scored as separated. Therefore, a meiotic figure like this is considered to have only three arm probe spots with one pair of sisters exhibiting premature loss of arm cohesion.

Premature loss of arm cohesion for both sets of sister chromatids results in four arm probe spots. In all four examples shown, centromeric cohesion is intact. Once mastered, rolling the oocytes for one genotype can be done in approximately 25 minutes if it is performed properly.

After watching this video, you should have a good understanding of how to roll oocytes, perform FISH using arm probes and score images to determine the state of meiotic arm cohesion in drosophila oocytes. While performing the hybridization and washes, it is important to remember to be patient. Not waiting long enough for oocytes to settle after each wash step will reduce the yield of oocytes for imaging at the end.

It may be possible to optimize this protocol to combine FISH with amino staining in order to answer additional questions about protein localization or the structure of the meiotic spindle while also monitoring the state of arm cohesion. The development of this technique will pave the way for researchers in the field of drosophila meiosis to explore the mechanisms that ensure and disrupt meiotic arm cohesion and impact chromosome segregation.