Trays of embryos are collected at 12 hour intervals and stored in the cold room in a covered container.  Stack the trays in an alternating fashion so each tray doesn't crush the embryos on the tray below.  The trays can be stored for up to 3 days - don't use embryos that have been stored for longer.  Trays from 3 cages collected over 3 days typically yield about 100 grams of embryos.

Combine all the ingredients in a volume of 800 ml and stir until everything has dissolved.  Add water to bring the final volume to 1 liter and store at 4 ^oC.

Just before starting the preparation of extract, add the following.
 

Final concentration	Addition per liter of Buffer I
1 mM DTT	1 ml of 1M (stock stored in freezer)
1 mM Na Bisulfite	0.19 g
0.2 mM PMSF	1 ml of 0.2 M dissolved in Ethanol

Combine all the ingredients in a volume of 200 ml and stir until everything is dissolved.  Add water to bring the final volume to 250 ml and store at 4 ^oC.

Just before use, add the following.
 

Final concentrations	Additions per 250 ml of Buffer AB
1 mM DTT	0.25 ml of 1 M
1 mM Na Bisulfite	0.048 g
0.2 mM PMSF	0.25 ml of 0.2 M dissolved in Ethanol

Combine all the ingredients in a volume of 1.5 liters and stir until everything is dissolved.  Add water to bring the final volume to 2 liters and store at 4 ^oC.

Just before use, add the following.
 

Final concentrations	Additions per 2 liters of Buffer C
1 mM DTT	2 ml of 1 M
0.5 mM Na Bisulfite	0.19 g
0.1 mM PMSF	1 ml of 0.2 M dissolved in Ethanol

• Prechilled ultracentrifuge, chilled Ti70 rotor and 6 polycarbonate ultracentrifuge tubes with red caps, black plugs and rubber sealing rings.  Some ultracentrifuges don't begin chilling until a vacuum has formed.  Familiarize yourself with the centrifuge so you can be certain it is chilled before use.  You might have to do a low speed spin with only the rotor to have the refrigeration turn on.
• Prechilled sorvall centrifuge, chilled Superlite GSA (Model SLA-1500) rotor and four 250 ml centrifuge bottles, chilled SS34 rotor and six 40 ml centrifuge tubes.
• Yamato homogenizer located in the cold room.
• Three funnels each containing a sheet of miracloth folded to line the inside of the funnel.  Rinse the miracloth with water and allow it to drain.  Support the funnels over 500 ml flasks and cover the miracloth with plastic until use.  Prechill everything in the cold room.
• Paint brush, screens and holders for collecting embryos.
• Rotating wheel placed in the cold room for mixing lysed embryos.
• Be certain to have at least 250 ml of bleach available.

1. Embryos that have been laid on grape plates for 12 hour periods are generally used. The plates are transferred every 12 hours to the cold room where the embryos can be stored for up to three days. While in the cold room, the trays of embryos are kept in a large plastic container covered with a lid so that they do not dehydrate.

 
2. Collection of the embryos is done by setting up a collection apparatus with a course Nitex screen to remove adults and a fine Nitex screen to trap embryos. The collection apparatus consists of three gray interlocking pieces of plastic pipe that support the screens. Check with someone who is familiar with the screens so you can be certain you are using the correct ones. One collection apparatus will usually handle the eggs from 18 to 20 grape plates. I prefer to set up two collection apparatuses and alternate between the two as I add suspensions of embryos so there is time for the liquid to drain.

 
3. Set up the collection apparatus in the large sink in the fly room where there is plenty of room to accommodate the grape plates. The embryos on the trays are wetted with water and then brushed into a plastic tub. Squirt water on the trays to rinse remaining eggs into the plastic tub. Try to avoid the strip of yeast paste. Empty the container of embryos into the collection apparatus after every four trays. You want to avoid having the eggs become anoxic. Throughout the collection period, intermittently splash tap water on the embryos in the collection apparatus. Be certain that the tap water is room temperature or less. Occasionally the tap water is warm in the summer. If this is the case, you will need to chill large flasks of water instead of using water directly from the tap.

 
4. Set the grape plates aside in a large covered container. If they have not dried and cracked, wash them off for reuse when you have time later in the preparation. It is a lot easier to reuse the grape plates than to pour fresh ones. (Clean the trays at some point when you have a break in the procedure. To clean, scrape most of the residual yeast off the surface of the agar with a spatula. Under running water, rub the surfaces free of eggs, yeast and fly debris using gloved hands or a sponge. Store the washed trays in a covered container in the cold room. If the trays are dried and cracked, they should be discarded in a plastic trash bag and the top of the bag should be taped shut.)

 
5. After all the eggs have been transferred from the trays, rinse the eggs remaining on the course screen to the fine screen below by squirting them with water. Remove the course screen and rinse the eggs with cold tap water. Carefully remove the upper pipe. Fold the screen in half so that it surrounds the eggs and gently squeeze the water out of the patty of embryos. Squeeze the folded screen and patty of embryos between several paper towels until the most of the water is out. Weigh the screen and patty. The screen weighs about 2 grams; the rest is embryos. Place the patty of eggs on ice.

1. Assemble two embryo collection apparatuses in the sink near the cold cabinet.

 
2. Prepare 500 ml of 1/2 strength bleach and put it in a 1 liter beaker with a stir bar on a stirrer.

 
3. Add 100 to 150 grams of embryos and stir the embryos for 90 seconds.

 
4. Pour equal amounts of the embryos into the two collection apparatuses.

 
5. Rinse each batch with approximately 1 liter of 0.7% NaCl/0.04% Triton X-100.

 
6. Extensively wash the embryos with tap water.  There is a hose attached to one of the taps behind the sink.  I begin with a slow flow and pinch the tubing so a fine jet stream of water can be sprayed on the embryos.  I increase the flow of the water once it begins to drain freely and continue to pinch the tubing so a forceful stream of water is applies to the embryos.  Rinse until all the Triton X- bubbles are removed and there is no scent of bleach.

 
7. Wash each paddy of embryos with 1 liter of distilled/deionized water.

 
8. Blot the embryos dry with a paper towel, weigh the embryos and place the paddies on ice.

1. Add DTT, PMSF and Na Bisulfite to buffer I and to buffer AB - don't forget!!!!

 
2. Suspend the embryos in 3 ml of Buffer I per gram of embryos.  Stir with a spatula over a period of minutes so the large chunks of embryos are dispersed.

 
3. Place a beaker under the Yamato homogenizer to catch solutions.  Wet the Yamato homogenizer with a small amount of Buffer I and then start it turning at 1000 rpm.  Pass the slurry of embryos through the device and collect the lysate in the beaker.  You may need to interrupt the process to transfer some of the lysate to another beaker if the beaker becomes too full.

 
4. After completing the first pass, pass the embryo lysate through the homogenizer a second time.

 
5. Pour lysate from 30 to 40 grams of embryos onto a single miracloth filter which is mounted in a funnel.  For 100 grams of embryos, you will need 3 filters set up.  After most of the lysate has drained through the miracloth, rinse the retentate with an additional 3 ml of Buffer I per gram of embryos.

 
6. For 100 grams of embryos, you should have approximately 600 ml of lysate.  Distribute this evenly among three 200 ml centrifuge tubes.  Centrifuge the material in a precooled Superlite GSA rotor at 9K for 15 minutes.

 
7. Slowly pour off the supernatants and resuspend the nuclei in 1 ml of Buffer AB per gram of embryos.  The buffer AB should already contain DTT and PMSF (see above).

 
8. Resuspend the nuclei using a glass dounce and a B pestle.

 
9. Measure the volume of resuspended nuclei and distribute 20 to 25 ml portions equally among polycarbonate ultracentrifuge tubes.

 
10. Add 1/10 volume of 4 M ammonium sulfate pH 7.9 (room temperature) to each tube.  The 4 M ammonium sulfate is added quickly to an individual tube, the tube is capped and then mixed rapidly by inverting the tube.  This is repeated for each of the tubes.  The solution should become very viscous.  If it doesn't, something is wrong.

 
11. Mount the tubes on the rotating wheel in the cold room and mix the tubes end over end for 15 to 20 minutes.

 
12. Spin tubes in a pre-cooled Ti70 rotor at 35 K for 1 hour (be certain the ultracentrifuge is also prechilled).  One hour is the time the samples spend at 35K.  It usually takes 5 to 10 minutes for the vacuum to form and for the rotor to reach maximum speed.

 
13. Immediately after the spin is completed, carefully remove the tubes from the rotor.  There will be a gelatinous pellet at the bottom and a white cloudy layer at the top.  In the middle will be a clear brownish yellow fluid.  Collect this layer with a 10 ml pipette by plunging the tip of the pipette well below the white layer and sucking steadily.  Leave behind the bulk of the lipid layer (a few ml).  Collect material from all the tubes in one graduated cylinder and determine the total volume of extract.

 
14. Place a beaker with a stir bar in a tray of ice and place the tray on a stirrer.  Add the lysate and start the stirrer.  Steadily add 0.3 g of finely ground ammonium sulfate per ml of lysate over a period of 5 minutes.  The solution should be stirring.  Leave the solution stirring for an additional 10 minutes.

 
15. Pour the lysate into 40 ml sorvall tubes and centrifuge the material at 15 K for 20 minutes in a pre-cooled SS34 rotor.

 
16. Pour off supernatant, drain well and then dry the sides of the tubes with a Kimwipe.  These pellets can be stored on ice in the cold room for up to 1 week but I usually don't leave them for more than a day or two.  Cover the tubes with parafilm during storage.

 
17. Add PMSF, DTT and Na bisulfite to 2 liters of cold Buffer C and resuspend pellets by adding 0.2 ml of Buffer C/gram of embryos. Disperse the pellet using the pestle from a dounce.  Fully resuspend the pellet using the dounce.

 
18. Dialyze the extract in the remaining 2 liters of buffer C.  The lysate is dialyzed in Spectra/Por Membrane 1 (cut-off: 6-8000).  One end of the tubing is folded over and sealed with an orange clip.  The tubing is rinsed several times with distilled/deionized water and once with a small amount of buffer C.  Add the lysate, fold over the open end and seal with an orange clip.  Place the dialysis bag in the beaker of buffer C and set the solution stirring in the cold room.
    
    
19. Dialyze the extract until the conductivity is equal to that of between 0.12 and 0.15 M HEMG.  Begin checking the conductivity after about 2 hours of dialysis.  Remember to mix the contents of the dialysis bag before removing a small portion of extract for each measurement.  I usually dilute 10 ul of extract in 20 ml of water to measure the conductivity and compare this to the conductivity of 0.1 M HEMG that has been diluted in 20 ml of water.  The dialysis usually takes from 2 to 4 hours.
• The original Biggin protocol indicated that the samples should be dialyzed until they reach a conductivity of 0.1 M HEMG.  A lot of protein precipitates as the extract approaches 0.1 M and below so I have tried to reduce what might be lost by stopping the dialysis at a slightly higher salt concentration.
• You may see an increase in protein yeild if you mix the contents of the dialysis bag every 20 to 30 minutes.  This manuever was recommended by David Price and makes sense to reduce formation of localized regions of low salt where proteins may be prone to precipitation.
• HEMG corresponds to 25 mM HEPES pH 7.6, 12.5 mM MgCl2, 0.1 mM EDTA pH 7.9, 10% glycerol.  The number preceding HEMG corresponds to the concentration of KCl.  For example, 0.1 M HEMG contains 0.1 m KCl.  Buffer C is simply 0.04 M HEMG.
• Don't forget the DTT and protease inhibitors.
20. Once the dialysis is complete, use a pipette to transfer the extract to clean tubes.  Spin out any precipitated protein by spinning the extract for 20 seconds in a cold microfuge or for 5 minutes at 9K in a Sorvall SS-34 rotor.  Transfer the clear supernant to new tubes and discard the precipitate.

 
21. Aliquot as desired, freeze the extract in liquid nitrogen and store at -70^oC.  Reserve at least on small aliquot of extract to determine the protein concentration.  Biggin reports that approximately 5 mg of protein are obtained per gram of dechorionated embryos.