University of Kentucky
1. Digest good-quality genomic or plasmid DNA to completion with restriction enzymes. Use AT LEAST 2 micrograms of genomic DNA per lane; 5 micrograms per lane is ideal. Less plasmid DNA is needed (250 ng-1 microgram). If more than one DNA sample is to be compared, you should load equal quantities of each. Digestion should proceed for at least 1 hour, and usually for no more than 3-4 hours.
2. Make an agarose gel. Use 1X TAE, and pour the gel no more than 1/4 inch thick. Add loading dye to digests (1 microliter of dye per 3 microliters of sample). Load samples into wells, and electrophorese relatively slowly until the dye passes off the end of the gel. For an 11 X 14 gel, 30 volts for 6 hours usually gives good results. (Running the gel slowly results in thinner, sharper bands on the Southern).
3. Ethidium-stain and photograph the gel (ethidium bromide is a carcinogen; stained gels should be handled only with gloved hands, and as little as possible: in an emergency ethidium can be removed from the skin with 95% ethanol). The stained gel should be placed in a glass cake dish, and shaken gently in the following series of solutions for the specified times (see end of protocol for recipes):
1. Depurination solution: 15 minutes
2. Denaturation solution: 30 minutes
3. Rinse briefly in sterile distilled water.
4. Neutralizing solution: 2 X 15 minutes
5. 20X SSC: 2 X 15 minutes
4. Prepare turboblotter as specified in instructions printed on blotter card. Have ready a clean dish containing sterile distilled water and a clean dish containing 20X SSC. The dry filter can be labeled with a soft lead pencil. Presoak the labeled nylon filter for about 5 minutes in the water, then transfer to the 20X SSC (filters should be handled at the edges only with clean forceps or gloved hands). Soak the appropriate blotter papers in 20X SSC, and put the blotting stack together in the order specified. Be sure to eliminate any air bubbles between the gel and the nylon filter. If the gel is smaller than 11 X 14, use parafilm to mask the edges of the blotter papers and the membrane underneath the gel to prevent the blotting solution from by-passing the gel. When the stack is complete, load the turboblotter with 20X SSC, add the pre-soaked wick paper, and the blotter card. Leave the blot for at least 4 hours (can be left overnight, if necessary).
5. Disassemble the turboblotter stack. Discard blotting papers in the regular trash, and the gel in the ethidium waste container. Wash the turboblotter apparatus thoroughly in plain water. Place the blot inside the Stratalinker on a clean paper towel. Press the auto-crosslink button, then the start button. When the cross-linking is complete, allow the blot to air dry on the bench for about 1 hour, then seal it in a heat-seal bag until use.
Depurination Solution: 20.8 mls of concentrated HCl in 1 L of H20 (use ultrapure water). Be careful with the concentrated acid: wear lab coat, gloves, and safety glasses, pour and mix in the fume hood. Add the acid to the water carefully, it will steam a bit.
Denaturing Solution: 20 g of NaOH, 58.4 g of NaCl per liter of ultrapure water. NaOH is caustic, so be careful with it.
Neutralizing Solution: 78.8 g Tris base, 87.6 g NaCL in 800 mls of ultrapure water. Adjust the pH to 7.4. Add ultrapure water to 1 liter.
20X SSC: 175.3 g NaCl, 88.2 g of trisodium citrate (citric acid); add ultrapure water to 1 liter.
1. Labeling the Probe using the Genius System: Labeling works much better on linear DNA, so plasmids should be linearized first and then purified from an agarose gel using the Quiaex Kit. Before beginning, thaw Genius tubes 5 and 6 on ice. To label the fragment, put 15 microliters of the Quiaex DNA (500ng-1microgram) into a microfuge tube, and place it in the boiling rack in a boiling water bath for 10 minutes. The tube should then be placed immediately on ice. After 2 or 3 minutes, add 2 microliters of hexanucleotide mix (Genius tube #5), 2 microliters of dNTP mix (tube #6) and 1 microliter of Klenow enzyme (tube #7). Mix together, centrifuge briefly to collect at the bottom of the tube, then incubate the tube overnight at 37 C. After incubation, add 1 microliter of 0.5M EDTA to the tube to stop the reaction. Then add 10 microliters of 7.5 M ammonium acetate and 60 microliters of chilled 95% ethanol. Mix together and incubate at -20 C for 2 hours. Spin in the cold microfuge at top speed for 15 minutes. Wash the pellet one time with cold 70% ethanol, and resuspend in 50 microliters of TE buffer. The labeled probe can be stored at either 4 C or -20 C.
2. Hybridization in the Rotisserie Oven: Turn on the oven and set to 65 C. Place empty bottles in the holder and start the rotisserie so that the bottles can pre-warm. Turn on the water bath and set to 65 C. Make up 20 mls of hybridization solution for each blot (see the end of the protocol for recipe). Also make about 50 mls of 2X SSC for each blot. Pre-warm both of these solutions in the water bath. Loosly roll up the blot along with a screen and place the roll in the warmed bottle. Add warmed 2X SSC to the bottle, gently unroll the blot and pour out the SSC. Add 10 mls of the hybridization solution and place in the rotisserie to pre-hyb the blot for at least 1 hour (blot can pre-hyb longer if necessary). After prehybridization is complete, boil 10-20 microliters of the labeled probe mixed with 90-80 microliters of SDW (total of 100 microliters) for 10 minutes in a boiling water bath. Place immediately on ice. After the probe cools, add it to the remaining 10 mls of pre-warmed hybridization solution. Pour out the pre-hyb solution and add the hybridization solution containing the label. Return the bottle to the oven and incubate for at least 16 hours (overnight). If you are re-using a probe that is already in hybridization solution (see below), heat the tube containing the mix to 95 C for 10 minutes in a (not quite) boiling water bath, and then add the mix to the bottle containing the blot. Blot can hybridize longer if necessary. The rotisserie should be set at a speed of 7.
3. Rinsing the Blots: After hybridization is complete, make up rinsing solutions as follows:
solution 1: 100 mls of 2X SSC/0.1% SDS
solution 2: 100 mls of 0.1X SSC/0.1% SDS (pre-warm to 65 C for a high stringency blot).
Pour off the hybridization solution. This can be placed in a plastic test tube, labeled with the date and the name of the probe, and stored at -20 C to be used again (see above). Remove the blot gently from the bottle and place it and the screen in a clean glass cake dish. Return the bottle to the oven to stay warm. Wash the blot twice in 50 mls of solution 1, 5 minutes each wash, at room temperature. Roll the blot and the screen up again and return to the warmed bottle. Add 50 mls of pre-warmed solution 2, unroll the blot, and place the bottle in the oven for 15 minutes. The rotisserie should be turning at top speed. Repeat this rinse once. For a lower stringency blot, the temperature of this second rinse can be lowered. The exact temperature must be determined empirically.
Standard Hybridization Solution (40 mls)
10 mls 20X SSC
400 microliters 10% (w/v) N-lauroylsarcosine (filter sterilized)
40 microliters 20% (w/v) SDS (filter-sterilized)
8 mls Genius stock blocking reagent (see below)
21.56 mls sterile ultrapure water
Stock Blocking Reagent: Dissolve blocking reagent in maleic acid buffer (see below) to a final concentration of 10% (w/v) with shaking and heating. DO NOT BOIL. Solution will be cloudy. Autoclave and store at 4 C.
Maleic Acid Buffer
0.1M Maleic Acid
0.15 M NaCl
Adjust pH to 7.5 with concentrated or solid NaOH: autoclave.
1. Equilibrate the washed blot in maleic acid buffer (see end of protocol for recipes). Do all of the following washes in clean plastic boxes on the shaker at room temperature.
2. Wash the blot for 1 hour in blocking buffer.
3. Mix up the antibody solution: Add 2 microliters of antibody (Genius tube #8, stored at 4 C) to 20 mls of blocking buffer. DO NOT CONTAMINATE THE ANTIBODY: PIPETTE GENTLY TO AVOID FOAMING. Place the blot in a clean plastic dish, add the antibody solution, and cover the blot with saran wrap, excluding any air pockets. Shake gently for 30 minutes at room temperature.
4. Rinse the blot twice for 15 minutes each with excess maleic acid buffer (50-100 mls).
5. Rinse the blot for two minutes in 20-50 mls of detection buffer.
6. Place the blot into an opened out heat-seal bag. Pipette 2 mls of CSPD solution into the bag and fold the bag over the blot, allowing the solution to cover the blot completely. Secure the bag with paper clips and place at 37 C for 15 minutes.
7. Seal the blot in the heat-seal bag. Place the blot in a developing cassette, add a sheet of film in the dark room (DO NOT EXPOSE X-RAY FILM TO LIGHT). Allow to expose at room temperature for about 1 hour. Develop the film using the automatic developer in the agronomy darkroom downstairs. The blot can be stored in the refrigerator and re-probed at a later date. If the signal is weak, the blot can be incubated ON at 37 C and then re-exposed for one hour the next day. However, there is no point in incubating it any longer than overnight. (That's all you get!! :)
Maleic Acid Buffer: (see Southern Blot: Part 2)
1 part Genius stock blocking reagent
9 parts Maleic Acid Buffer
100 mM Tris base, pH 9.5
100 mM NaCl
50 mM MgCl2
Prepare detection buffer from sterile stock solutions to avoid precipitation of MgCl2. Adjust the pH to 9.5 prior to addition of the MgCl2 or it will precipitate.
Wear gloves. Add 20 microliters of CSPD to 2 mls of detection buffer. Make immediately before use, and do not make more than you need. DO NOT CONTAMINATE CSPD.
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