4.7 Miniprotoplast preparation for BAC library

Naomi Sumikawa and Takashi Murata



Protonemata were cultured on BCDAT medium layered with cellophane for 6 days under continuous light after inoculation, and then they were kept in the dark for 18-24 hrs. The dark incubation removes starch grains from chloroplasts. Do not use BCDTAG medium, which contains glucose, because glucose in the medium prevents removal of starch. It is very important to keep more than 18 hours in the dark in order to isolate protoplast from debris (see Fig. 1).



Solutions and tubes needed

1. Digestive solution: 2% driselase and 10% mannitol in H2O (160 ml for 4 tubes) at room temperature


2. 10% mannitol in H20 (kept in ice-water)


3. MP solution          50 ml

    PS/HEPES (pH 7.3)        18 ml

    0.6 M Sorbitol     30 ml

    1 M MgCl2            2 ml

    Freshly prepare MP solution for each experiment.


   PS/HEPES (pH 7.3)

    Percoll (Amersham)       1 L

    sucrose         250 g

    Dissolve and adjust pH to 7.3 with HEPES powder.

    Store at 4C


3. 35 ml polycarbonate cfg tubes (kept in ice-water or at 4C°).


4. One 50 ml polypropylene tube containing 10 ml 0.55 M sucrose in H2O. Keep in ice-water.


5. It is preferable to keep all plastic, glass dishes, and solutions except the digestive solution and the MP solution at 4C. This may not be seriously necessary.


Keep the digestive solution and the MP solution at room temperature.


6. Use wide-pore round-tip pipettes kept at 4C°.





0. Chill cfg and swing rotors (for 35 ml and 50 ml) at 2C°


1. Harvest eighty 9-cm petri dishes of protonemata layered with cellophane.

You can get 6.4 x 10^7 miniprotoplasts. If you need, you can reduce the scale.


2. Move protonemata from 20 dishes to a 35 ml round-bottom polycarbonate cfg tube (PC tube) containing 40 ml digestive solution [2% driselase and 10% mannitol in H2O]. Four tubes for 80 dishes.


3. Incubate for 40 min at 25. Gently but thoroughly rotate tubes once in 10 min.


4. Filtrate through a nylon seat with 50 µm mesh to new four PC tubes. You can use a nylon seat for four tubes. If flow-through becomes worse, wash the seat with 10% mannitol.


5. Store for 5 min at room temperature for complete digestion.


6. cfg. 1,000 rpm / 170 x g, swing rotor, 2 min at 2C°


7. Discard sup with wide-pore pipette (NO Decantation) and keep on ice.


8. Gently add 5 ml ice-cold 10% mannitol to each tube and gently suspend with wide and round-pore pipette on ice.


9. Add 25 ml of ice-cold 10% mannitol and gently mix.


10. cfg. 1,000 rpm / 170 x g, swing rotor, 2 min at 2C°


11. Repeat 7, 8


12. Gently layer 5 ml x 4 suspended solution on 0.55 M sucrose solution in a ice-cold 50 ml polypropylene tube.


13. cfg. 2,000 rpm / 670 x g, swing rotor, 10 min at 2C°


14. Collect middle green zone (Fig. 1a) with cold pipette and gently pour to two new 35 ml PC tubes on ice.


15. Slowly add 30 ml ice-cold 10% mannitol with gentle mixing on ice.


16. cfg. 1,000 rpm / 170 x g, swing rotor, 2 min at 2C°


17. Discard sup with wide-pore pipet (NO Decantation) and keep at room temperature.


18. Gently add 25 ml MP solution and gently and thoroughly mix.


19. cfg. 15,000 rpm / 25,000 x g, angle rotor, 45 min at 20C°


20. Chill a 35 ml PC tube on ice.


20. Move a lower zone (Fig. 2) to the ice-cold PC tube.


21. Slowly add 30 ml ice-cold 10% mannitol with gentle mixing.


22. cfg. 1,000 rpm / 170 x g, swing rotor, 2 min at 2C°


23. Discard sup with a wide-pore pipette (NO Decantation).


24. Add an appropriate solution to suspend ppt for following experiments.