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Early Cold Shock Gynogenesis
(Manda Cox, Grainger Lab, 03/2004; updated 4/2007)

Day 1

  1. Females of interest are gathered.

    •  There is no real limitation to the amount of females that can be screened in a given day, other than the time constraints of a normal workday. We have successfully handled up to 15 females in one day using two workers. The problem lies in scoring that amount of subsequent embryos, which takes considerably longer. On average, 4-6 frogs can be easily handled in one session and successfully scored.

  2. Females are primed via injection with an insulin syringe with 10 U of human chorionic gonadotropin (HCG) diluted to 100 uL total volume with the supplied diluent (phosphate buffered saline).

    •  For details on HCG injection please see that section on our website.

  3. Females can be returned to a 25 degree incubator overnight or left out at room temperature.

Day 2

  1. Boost females with 100 U of HCG in 100 uL total volume.

    •  The females should begin ovulation in 3-4 hours. At least 8 hours should be allowed between the prime and boost, with no longer than 72 hours. Improper injection technique is one of the top reasons for not obtaining eggs later in the experiment.

  2. Make following solutions (for instructions, see end of protocol):

     0.05X MBS + 0.1% BSA (1L)

     0.1X MBS (1L)

     0.1X MBS + gentamicin (1L)

     1X MBS + 0.1% BSA (50 mL)

  3. Fill 2-50mL conical tubes with part of the previously made 0.05X MBS + 0.1% BSA and pre-chill in a slushy ice water bath, generally kept in a 2 or 4 degree Celsius refrigerator. At the same time, pre-chill your vessel for placing embryo plates in the ice water bath.

    •  The 0.05X MBS + 0.1% BSA will be used to apply the cold shock. The ice water bath should be 1-3 degrees Celsius and the tubes fully submerged. You should pre-chill what you've already made to avoid the effects of pH or concentration gradient on the embryos. You should always chill more than you anticipate using in the event the tube is spilled, thus ruining your experiment for the next three months. Two tubes should cover 4-5 frogs easily. If more frogs than that are used, you should chill more tubes. An appropriate water bath can be made using a laboratory style bucket with ice and water. A suitable vessel for placing embryo dishes in is the lid of a P-1000 tip box. It is important to remember to chill the lid to avoid thermal variation or heat transfer during the cold shocking.

  4. Sacrifice 2-3 males for making the sperm slurry used. Place the males in a MS222 bath (temperature = 20-25 degrees Celsius).

    •  This should be done at most 30-45 minutes before ovulation. 2 males can comfortably be used for 4-5 females. More females than that will require more males. A general rule is 2 females/1 male. It will depend upon how you aliquot the sperm slurry later.

  5. While the males are being euthanized, place 2 mL of 1X MBS+ 0.1% BSA in a 15mL conical tube (if using 2 males). Place 100 uL of 1X MBS + 0.1% BSA in a microfuge tube for each of the males dissected.

  6. Glaze 60 x 15 mm petri dishes with 1X MBS + 0.1% BSA. You will need 3 petri dishes/ female tested.

    •  The easiest way to glaze the dishes and not waste solution is to pour some 1X MBS + 0.1% BSA into one or two dishes, then transfer it to each of the remaining dishes. Glazing is essential, so that the sperm will not stick to one place in the dish and instead, will move fluidly around the dish. The three dishes will correspond to the wild-type outcross control dish (used as a control to determine the level of endogenous background and will serve as a way of propagating the phenotype), the haploid control dish (used to determine if sperm irradiation was successful and can be scored for certain types of phenotypes) and the cold shock dish (experimental dish that will contain the gynogenotes).

  7. Dissect the testes from the male. Roll the testes around on a paper towel to remove excess blood and fat from testes. Place 1 pair of testes in each of the microfuge tubes prepared in step 5. Using a sterile micropestle, homogenize the testes in each of the tubes. Consolidate all the homogenized testes into one of the existing microfuge tubes and rinse the micropestle with 1 mL of 1X MBS+ 0.1% BSA, collecting the rinsing in the microfuge tube containing the consolidated sperm. Transfer the entire contents of the sperm prep to the 15 mL conical and allow the contents to settle for about 1 minute.

    •  When dissecting be sure not to tear the testes, as sperm will leak out resulting in a lower fertilization. Allowing the sperm to settle is essential because small chunks of tissue will remain and should be used only for the wild-type outcross control dish. Tissue in the haploid or cold shock dish will shelter individual sperm cells and they will escape irradiation.

  8. Aliquot the sperm slurry into the glazed 60 x 15 mm petri dishes. The cold shock dishes should be aliquoted first (see note above), followed by the haploid dishes and finally the wild type outcross control dishes. Typically, 300 uL of slurry are pipetted into the cold shock dishes, 200 uL into the haploid, and 200-250 uL into the WT outcross dishes.

    •  The irradiation process will kill off some sperm cells, thus why more are added to the cold shock dishes. The haploid phenotype is also dictated by genetics; many embryos will be completely unscorable, while others will be perfect haploids. Therefore, if you do not have enough sperm, it's always best to eliminate several of the haploid dishes. If it is imperative to have haploid dishes for all the females, you will need more males for the experiment—calculate based on the above volumes. Decreasing the aliquot volume tends to lead to decreased fertilization in the dishes and, therefore, is not always the best option for “stretching” your sperm slurry.

  9. Irradiate the haploid and cold shock dishes with UV light. Be sure to remove the lids, as the UV light cannot penetrate the lids of the petri dishes. Using our Fisher Biotech XL crosslinker, the sperm is irradiated at 1000 x 100 uJ/cm2.

  10. Immediately, but gently, squeeze the females eggs DRY into individual 100 x 15 mm petri dishes. Once eggs are collected, aliquot the eggs into the experimental dishes containing sperm using a disposable pipet glazed with 1X MBS + 0.1% BSA. More eggs should always go into the cold shock dish (2-3X the WT control dish). The haploid dish should, generally, receive the least amount of eggs.

  11. Swirl the dishes to ensure mixing of the sperm and eggs and let sit for 2-5 minutes. Flood dishes with 0.05X MBS + 0.1% BSA and let incubate at ambient temperature for EXACTLY 5 minutes.

    •  Approximately 30 seconds before time is up, you can start removing the media from the embryos to get ready for the application of the cold media. During the incubation time, be sure that your pre-chilled media temperature is between 1-3 degree Celsius. If it is not, you can generally warm it up by taking it out of submersion and placing it in the P-1000 lid that should be in the ice water bath; this allows for a small amount of warming. The experiment will be the most successful if the cold media is applied at exactly 5 minutes, rather than taking the 30 seconds to a minute to remove the RT media then apply the cold media once time has expired.

  12. Remove the RT media from the cold shock dishes only and replace with the cold media. Stack the dishes in the lid of the P-1000 box and incubate in the refrigerated ice water bath for EXACTLY 10 minutes.

    •  Do not stack the dishes over two high in the box or latent heat transfer will occur.

  13. Remove the cold media from the dishes and replace with the RT 0.05X MBS + 0.1% BSA that you previously used. Keep at ambient temperature until cleavage begins, approximately 1-1.5 hours later.

    •  It is imperative that the cold media is left on the entire 10 minutes and not removed before. Removing the media at this stage should be easier because the eggs have developed a thick jelly coat and will usually stick to the dishes. At the end of 10 minutes, you should remove the cold media from all the dishes first, then go back and add the RT media to them all. This will be henceforth called the “assembly line” method. Do not remove cold media from one and add the warm media, then move onto the next dish.

  14. When the eggs begin to cleave, dejelly them using 2% cysteine (in 0.1X MBS) for 6 minutes, while swirling the dishes.

    •  Since there are so many dishes, it's easiest to swirl them using a rotating surface such as those used with hybridization or gel staining. If the eggs are not swirled, the cysteine will eat away the jelly coat unevenly and destroy the embryos.

  15. After dejellying the embryos, remove the cysteine from each of the dishes and rinse the embryos at minimum 4x using 0.05X MBS + 0.1% BSA using the “assembly line” method.

  16. Sort the embryos into 100 x 15 mm Petri dishes using a fire polished Pasteur pipet in groups no larger than 100 embryos/dish. The dishes should contain 0.1X MBS + gentamicin. The embryos are then kept at 22 degrees Celsius for the next several days.

  17. The females' water is changed and supplemented with 1g/L of sea salt as a prophylactic means of soothing the females.

Days 3, 4, and 5

Embryos are scored each day and their media changed to fresh 0.1X MBS + gentamicin. Dead embryos are removed each day. The day after cold shocking (day 3), the embryos should be looked at first thing in the morning. If a dish has a lot of debris from dead and decaying embryos, the remaining embryos are transferred to a new dish (especially on Day 3). Once the embryos begin swimming on their own (generally on Day 4), a disposable transfer pipet can be used to pick them up. Once the tadpoles can swim, they should be lowered to a dish density of no more than 50 per dish.

If the embryos are swimming around too much to be scored (generally day 5 +), the tadpoles may be anesthetized by adding 10-15 drops of 1:500 MS222 to the dish. Once the scoring is complete, the embryos must be transferred to fresh media to wake up.

Day 6

Embryos are transitioned to 0.1X MBS without gentamicin and are fed sera micron. The sera micron solution should have a very slight tint of green to prevent excess food and waste in the dishes. Feeding should roughly correspond to the development of the gills in the tadpoles.

Day 8

If you want to keep the tadpoles this long, they should be transferred to a larger volume container such as a beaker or mod system. If transferring to a beaker, dump the entire contents of each dish into the beaker and dilute 1:1 with dechlorinated water. Continue to feed each day. If placing the embryos in a mod system or equivalent, please refer to the husbandry section.



10% BSA (50 mL)

5 grams of Fraction V bovine serum albumin

Dilute to 50 mL with sterile water in a conical tube

Freeze at -20 degrees Celsius when not in use.

Helpful Hint: don't shake BSA (it foams terribly). Instead, add about 10 mL sterile water to tube, then add the 5 g BSA, then top off with more water. Place on a nutator and allow solution to nutate for 15 minutes to dissolve. Then bring up to volume if necessary.


0.05X MBS +0.1% BSA (pH 7.5 + 0.1; 1L)

5 mL 10X MBS (sterile) salts

10 mL of 10% BSA (thawed)

bring to 1L with sterile water

**Make fresh daily


0.1X MBS (pH 7.5 + 0.1; 1L)

10 mL of 10X MBS salts (sterile) in 1L sterile water.

**May be kept indefinitely

0.1X MBS + gentamicin (pH 7.5 + 0.1; 1L)

same as above, except add total of 50 mg of gentamicin sulfate.

**Gentamicin sulfate is an acidic drug—the initial pH of this solution will be lower than 0.1X MBS

**Leftover should be refrigerated at 2-4 degrees Celsius and not kept more than several days.

1X MBS + 0.1% BSA (pH 7.5 + 0.1; 50 mL)

5 mL 10X MBS salts

500 uL 10% BSA

Dilute to 50mL in a conical tube with sterile water

**Make fresh daily


2% cysteine (pH 7.9 + 0.1; 150 mL)

3.0 grams cysteine in 150 mL of 0.1X MBS

**Make in sterile beaker or container




For more information about this project please contact us.
Last update: Feb. 13, 2008