Southern Transfer to Nylon membranes
Southern transfer is the classic method used to move DNA (usually in the form of restriction fragments) from agarose gels to a solid support, e. g. nylon membrane. The DNA is bound to the membrane in the same position (relative to other fragments) as in the gel. The use of a membrane in hybridization experiments is far more convenient than trying to hybridize in a gel. In a Southern transfer the DNA is first denatured in the gel, then the gel is placed on a buffer-saturated spongey surface, the membrane positioned on top of the gel, and dry blotting papers are added on top of the membrane. The buffer moves by capillary action through the gel and into the papers above. The denatured DNA in the gel moves passively with the buffer and is stopped by the membrane. DNA is permanently bound to the membrane by baking or cross-linking with UV. This protocol is designed for transferring DNA for RFLP analysis. The optimum size range of DNA fragments detected this way is 2 - 25 kb.
- For each gel to be run, melt a 250 ml stock bottle of 0.8% 1X TA agarose in the microwave (weigh the bottle before heating and make up lost volume with dH2O after agarose is melted). Cool to 50 degrees C; or prepare 250 ml of 0.8% Tris-Acetate-agarose following the recipe given here under Solutions.
- Seal both ends of a gel casting tray with the labeling tape and place it on a level surface. When the agarose has cooled to 50 degrees C, pour 250 ml TA agarose into the gel casting tray without forming air bubbles. If you see air bubbles, move them to the edge or pop them with a pipet tip and place a comb in the first set of slots on the gel casting tray. Place a 30-35 tooth comb in the first set of slots in the gel cast. Let the agarose solidify at room temperature for at least 30 minutes.
- Remove the tape, then place the gel tray in the electrophoresis box. Orient the wells towards the negative end (black leads) and pour 1X TA buffer into the box until the gel is submerged. Pull the comb gently out of the gel. (If gels are prepared several hours in advance do not pull comb until you are ready to load.)
- Load the samples in the designated lane orders, changing pipetmen tips with each sample. Use lambda DNA combination marker heated to 50 degrees C to melt the ''sticky ends''. (For more about these molecular weight standard markers, refer to appendix)
- Circulate the buffer using a low speed pump, e. g. a DIAS pump, with the buffer flowing from the negative end (black lead) to the positive end (red lead) of the gel box.
- Connect the leads to the power supply, place the timer on hold and adjust the voltage to a constant 40-45 volts. Run the gel for 18-20 hours (for Screening blots) or 20-24 hours (for Parent & Family blots).
- To stain the gels, pour 800-1000 ml of dH2O into a gel staining box, add 15 祃 of 10 mg/ml ethidium bromide (EtBr), mix and immerse the gel in the staining box. The stain solution should cover the gel completely. Place a gel restrainer on the tray to hold the gel in place.
- Shake gels at room temperature on a platform shaker for 30 minutes. Drain the stain solution, rinse the gel in dH2O to remove excess EtBr and photograph the gel with UV illumination using the Fotodyne UV/MP-4 camera. Mark the ends of the gel on the UV box by making small cuts to define where the final cuts in the gel or Southern blot will be, (e. g. to separate two families run on the same gel). Typical exposure time is f 5.6 for 1 second.
- To denature the DNA in the gels, by shake gels slowly in a freshly prepared solution of 0.6M NaCl, 0.2N NaOH (for 1 liter: 200 ml 3M NaCl + 20 ml 10N NaOH brought to 1000 ml with dH2O) for 30 minutes. Check frequently to be certain the gels stay immersed in the denaturant.
A common problem is that if the gel surface remains dry, the DNA will not denature properly resulting in poor transfer. Often the gels can be kept submerged with a sunken clean glass test tube positioned on top of the gel. Rinse gels briefly in dH2O.
- To neutralize the gels, shake slowly in 1.5M NaCl, 0.5M Tris-HCl pH8.0 (for 1 liter: 500 ml of 3M NaCl + 500 ml of 1M Tris-HCl). Check frequently to be certain gels stay immersed.
- If using prepared Southern set-ups, remove the top blot blocks from each and replace them with ones freshly soaked in 10X SSC. If a new transfer set-up is needed, wet blot blocks one by one by soaking them briefly in 10X SSC and transferring them to a fresh blotting tray. Layer blot blocks to a thickness of about 1 1/2". When layering the blot blocks, gently roll a clean test tube across the top to remove air bubbles between blocks.
- Fill the trays with 10X SSC buffer up to the top of the stack (but do not immerse the top of the stack).
- Cut 3MM Whatman papers (or use pre-cut 3MM Whatman paper) to the size of the blot blocks, individually wet them in 10X SSC, and lay one on each stack of blot blocks. Remove air bubbles by rolling the test tube as above.
- Cut Zetabind (or any other nylon membrane used as a solid support) slightly larger than the gels (e. g. 8" x 8") with a pair of clean scissors (wear gloves when handling membranes and protect membranes by working on clean 3MM paper). Label the membranes across the top with the respective family/parent I.D. #, the enzyme and the blot number using a dry-erase marker and cut across the left hand top corner of the membranes for orientation purposes.
- Wet the nylon membranes briefly in dH2O, transfer to a tray containing 10X SSC and soak for 20 minutes.
- Trim the gels at the wells, the bottom and sides (if necessary) and slide onto the blotting tray. Remove any air bubbles from under the gels with gloved fingers.
- Place the correctly labeled membrane on top of each corresponding gel with the notch oriented towards the top left hand corner. Remove air bubbles. If more than one family is on a gel, mark the membrane edges where cuts will be made after the transfer is complete.
- For each gel, wet another piece of 3MM Whatman paper (cut to the size of the gel) in 10X SSC and place it on the membrane. Remove air bubbles. Cover the exposed portion of the wet blot blocks with strips of parafilm or Saran wrap (which act as a wicking barrier). Be certain not to cover any gel lanes containing DNA. Place a stack of dry blot blocks (~ 2 cm high) on top the 3MM Whatman paper and place a set of paper towels (1-2 inches high) on top.
- Place a gel restrainer and a weight of about 500 g on top of each stack of paper towels (a 500 ml bottle or flask of water is approximately 500 g). Refill the blotting trays with 10X SSC to within 1/2 inch from the top of the wet blot blocks and allow the transfer to go 6 hours to overnight. Check the level of 10X SSC occasionally.
Taking down the transfers:
- Discard the soaked upper blot blocks and paper towels and transfer the membranes to a tray containing 500 ml 2X SSC. Gently rub each membrane with a gloved hand to remove residual agarose. Save the lower blot blocks in the tray and cover them with Saran wrap for future use. Label the blotting tray with the date.
- Wash the blots at room temperature in 2X SSC twice, 15 minutes each wash.
- Air dry the blots between two pieces of 3MM Whatman paper for at least 1 hour. Bake the blots (between Whatman papers) at 80癈 in the vacuum oven for 1-2 hours.
- Wash the blots in a shaking waterbath at 65 degrees C for 30 minutes in 500 ml of 0.1X SSC, 0.5% SDS (2.5 ml of 20X SSC + 25 ml of 10% SDS brought to 500 ml with ddH2O). This step appears to reduce the non-specific background often seen in the first hybridization of blots.
- Store the blots wet as follows: place very wet in a blot bag or seal-a-meal bag and seal the bag with a T-bar heat sealer.
- Fill out a new blot record sheet for each blot, tape the gel picture on a sheet of paper with all the relevant gel conditions written down and file both sheets in the proper blot book File the blot in its respective file folder.
Pour 16 liters of ddH2O into a 20 L carboy. Slowly add 3 kg NaCl.
Mix well. When the NaCl has dissolved, adjust the volume to 17.1 liters
Prepare in the chemical fume hood, and wear gloves and goggles:
To a 4L carboy add 2L of ddH2O and slowly add 1600 g of NaOH pellets.
1M Tris-HCl, pH8.0:
Adjust the volume to 4L with ddH2O.
Caution: This is an exothermic reaction. The solution gets very hot!!
Pour 5 liters of ddH2O into a 20L carboy. Slowly add 2500 g of Trizma
base with a stir bar stirring vigorously and bring the volume to 18
liters with ddH2O. Mix overnight.
Add 1000 ml of concentrated HCl. Adjust the pH to 8.0 with more HCl
and bring the volume to 20.65 liters with dH2O.
0.8 % 1X TA agarose:
Weigh 2.0 g of agarose into each 500 ml bottle. Add 250 ml of 1X TA to
each. Weigh the bottles before heating and make up lost volume with
dH2O after dissolving the agarose. Swirl the bottles to mix the agarose
well and either autoclave the agarose solution for 10 minutes or heat
in a microwave oven on HIGH for 6-8 minutes. Keep bottle lids loose
when heating the solution. Once the agarose is completely in solution,
swirl each bottle gently to mix the agarose evenly. Place the bottles
in a 50 degrees C waterbath for 20-30 minutes before pouring the
Dissolve 100g sodium dodecyl sulfate (SDS) in 500 ml of ddH2O, adjust
volume to 1000 ml and store at room temperature. Wear a face mask while
weighing out SDS.
20X SSC (20 liters)
Dissolve 3504 g NaCl and 1760 g sodium citrate in 10 L ddH2O.
(Use a 20 L carboy.) Adjust the volume to 20 L with ddH2O, and the
pH to 7.4 with several drops of concentrated HCl.
Southern, E.M. (1975). "Detection of specific sequences among DNA fragments separated by gel electrophoresis." J. Mol. Biol., 98: 503.
Sambrook, J., Fritsch, E.F., and T. Maniatis. (1989). Molecular Cloning - A Laboratory Manual (second edition), Cold Spring Harbor Laboratory, Cold Spring Harbor, New York. p. 9.31-9.44.