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Chorion and Vitelline Membrane Mechanical Removal


Chorion and Vitelline Membrane Mechanical Removal: A Method to Prepare Drosophila Oocytes for Direct Observation



- Begin with mature oocytes from fixed ovaries in a saline solution. Mature oocytes bear eggshells consisting of an outer chorion and inner vitelline membrane.

To mechanically remove the chorion and vitelline membrane, place the oocytes between the frosted parts of two pre-treated slides. Move the top slide in straight back-and-forth motions to create friction that rolls the oocytes.

Now, move the top slide at a slight angle to roll the oocytes in another direction. Avoiding circular motion, increase this angle in short increments until the top slide moves perpendicular to the bottom slide. This movement mechanically removes the chorions and vitelline membranes, enabling access for probes or stains into the oocyte.

Oocytes without chorions will appear long and thin, and those without vitelline membranes will have pointed ends. To separate the cleared oocytes from the surrounding debris, transfer the sample mix to a tube, and allow the oocytes to settle to the bottom while the debris floats at the top and can be removed.

In this protocol, we will remove chorion and vitelline membranes from mature Drosophila oocytes.

- To separate the late-stage oocytes, first, add 1 milliliter of PBSBTx to a shallow dissecting dish. Then, use a P200 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-microliter 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 with 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 P200 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 7 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 at 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 1 milliliter PBSBTx to a 15-milliliter conical tube. Swirl the liquid to coat the sides of the tube. Using a PBSBTx-coated P1000 pipette tip, transfer the rolled oocytes from the deep-well dish to the conical tube containing 1 milliliter of PBSBTx. Add an additional 2 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 P1000 to remove the top 2 milliliters of solution containing debris and discard.

After repeating the step for a total of three rounds of debris removal, use a PBSBTx-coated P1000 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.

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