Southern Blotting


* Gel box and combs
* Agarose
* 1X TBE
* Ruler
* 1.5M NaCl, 0.5M NaOH (in carboy in middle room)
* Blotting paper
* Hybond N+ transfer membrane
* 2X SSC (20X SSC in carboy in middle room)
* Whatman paper
* Pre-hybridization buffer (0.5M Na2HPO4, pH 7.2 / 7% SDS / 1mM EDTA)
* Template for probe labeling
* Random primer klenow labeling kit (available from Invitrogen or NEB)
* Film and cassette or phosphorimager cassette.



1. Prepare the appropriate amount and percentage (use the lowest percentage possible) of TBE-agarose gel. Let gel cool down.

2. Prepare gel box with combs. Make sure gel is level.

3. Add ethidium bromide (2.5ul of 10mg/ml EtBr stock in 100ml gel) to gel and pour gel.

4. Once gel is solid, cover it with 1X TBE until buffer is ~3 mm above the gel

5. Add loading buffer to each tube of sample DNA. Mix and spin down.

6. Load ladders and samples into the gel and apply voltage (100V is usually sufficient) until appropriate sizes are separated. Bands often look better if gel is run slowly.

7. Once the gel has run, photograph the gel with a ruler on the side, with the zero mark aligned with the inner side of the wells.

8. Place gel into 500 ml 1.5M NaCl, 0.5M NaOH (in carboy in middle room). Dye should change color after 15-20 minutes.

9. While gel is soaking, set up transfer apparatus with glass plate over a dish containing ~500 ml 1.5M NaCl, 0.5M NaOH. Pre-wet two pieces of whatman paper that are the width of gel and are long enough so that the ends are in the buffer and place on glass plate (make sure to remove any air bubbles). (picture 1A)

10. Cut a gel sized piece of transfer membrane (Hybond N+ or other appropriate product) and wet in 1.5M NaCl, 0.5M NaOH.

11. Place gel (upside down) on whatman paper and place membrane on top of gel (be sure to remove any air bubbles). Place two pieces of pre-wetted gel size whatman papers on top of membrane. (picture 1B)

12. Place 1.5 inch stack (1 package) of gel sized dry blotting paper on top. (picture 1C) Place second glass plate on top of blotting paper and weigh down with a 500 ml bottle of solution.

13. Let transfer overnight (at least 12 hours).

14. Disassemble transfer by removing blotting paper and turning the whole thing upside down.

15. Gel should now be on top of the membrane and should be covered by two pieces of whatman. Remove whatman without disturbing gel. Mark wells by using a VWR marker to poke through the wells so that the membrane is marked.

16. Remove gel and cut off the top left corner of the membrane.

17. Wrap membrane in saran wrap (do not rinse membrane before crosslinking).

18. Place in Strata Linker, and crosslink using the 'Autocrosslink' button.

19. Membrane is now ready for hybridization. Membrane may be stored for later use, wrapped in saran wrap at room temperature.

20. Place membrane in hybridization tube with DNA face away from the sides of the tube.

21. Add 25 ml pre-hybridization buffer pre-warmed to 65C (0.5M Na2HPO4, pH7.2 / 7% SDS / 1mM EDTA).

22. Incubate at 65C in hybridization oven with rotation.

23. While membrane is in pre-hyb, prepare labeled probe. For random primer klenow labeling, use kit from Invitrogen or NEB. Template should be at least 1kb (gel purified PCR fragments work well), although smaller fragments can work but you may need to optimize the hybridization temperature. When labeling is complete, purify on a G50 spin column. (if re-using old probe, simply thaw at 37C, denature 5 mins in boiling water bath and place on ice for 2 mins and then exchange with pre-hyb buffer).

24. Denature probe by placing at 95C for 5 minutes (make sure that you use an eppendorf caplock or place something heavy on the lid of the cap so that it doesn't pop open).

25. Place on ice for 2 minutes and then quick spin at room temp to collect droplets on side of tube.

26. Add probe directly to hybridization tube and leave overnight at 65C with rotation.

27. Remove hybridization buffer and either dispose of in liquid radioactive waste, or save for later use by storing in a 50ml falcon tube at -20C.

28. Wash membrane. The number and type of washes depends upon the sample and the probe.
Low stringency: 30 mins in 25ml 2X SSC, 0.1% SDS at room temp.
Medium stringency: 30 mins in 25ml 0.2X SSC, 0.1% SDS at room temp.
High stringency: 30 minsin 25ml 0.2X SSC, 0.1% SDS at 65C.
Washes should be done in hybridization tube on a rotator (room temp washes can be done with oven off). After each wash, before trying the next level of stringency, remove membrane from hybridization tube and check radioactivity with geiger counter (if you can find regions of the membrane that are hotter than others, ie bands, then stop washing. If there is no detectable radioactivity then the membrane was probably overwashed and the hybridization needs to be repeated. Dispose of wash solutions in liquid radioactive waste.

Alternate wash protocol:
Each of these washes should be performed with wash buffer that has been pre-warmed to 65C, then the wash is done either at room temp or 65C.
2x5 minutes in 40mM NaPi, 5% SDS, 0.1mM EDTA at room temp.
2x5 minutes in 40mM NaPi, 1% SDS, 0.1mM EDTA at room temp.
10-25 minutes in 40mM NaPi, 1% SDS, 0.1mM EDTA at 65C.
10-30 minutes in 0.2X SSC, 0.1% SDS at 65C.

29. Rinse membrane in 500 ml 2X SSC.

30. Remove membrane from rinse and allow to drip dry for 5-10 seconds.

31. Wrap membrane in saran wrap and expose to film or phosphorimager screen. Exposure generally takes 1-2 days on phosphorimager screen and about twice as long on film.

32. Membrane can be stored in this state for later use (see protocol for stripping membrane) or disposed of in radioactive waste when exposure is finished.