Localization of particular sequences within genomic DNA is usually accomplished by the transfer techniques described by Southern (1975). Genomic DNA is digested with one or more restriction enzymes, and the resulting fragments are separated according to size by electrophoresis through an agarose gel. The DNA is then denatured in situ and transferred from the gel to a solid support (usually nitrocellulose filter or nylon membrane). The relative positions of the DNA fragments are preserved during their transfer to the filter. The DNA attached to the filter is hybridized to radiolabeled DNA or RNA, and autoradiography is used to locate the positions of bands complementary to the probe. A sequence of 1000 bp that occurs only once in the mammalian genome can be detected in an overnight exposure if 10 µg of genomic DNA is transferred to the filter and hybridized to a probe several hundred nucleotides in length.
Complete digestion of genomic DNA is crucial but difficult. The average restriction fragment size varies according to the length of the recognition sequence, and also the GC content of the recognition site; AT rich sites occur more frequently. You don't want to see DNA hanging up at the mobility limit of the gel; you do want to see a few tight bands within the smear (these are repetitive sequences). To ensure complete digests, dilute the digest into a larger volume (thus diluting the enzyme inhibitors) and then precipitate and resuspend the cut DNA in the desired volume. You can try using 10 units of restriction enzyme per µg of DNA and digesting overnight (note: this is only useful for stable restriction enzymes- the Biolabs catalog has a table showing the stability of various enzymes). EcoRI is excellent at cutting genomic DNA, HindIII is good, others vary. After digestion, run an aliquot on an analytical gel to check the digestion: should see a smear with a bell-shaped distribution of intensity. EcoRV and XbaI are other 6-cutters with AT-rich recognition sites that work okay.
Cut off and autoclave pipette tips for handling high-molecular-weight genomic DNA.
Digest 10 µg genomic DNA in 400 µl total volume with 3 units restriction enzyme per µg DNA at 37°C for overnight.
(For random primer labeling you will also need to cut out the cDNA probe from vector.)
(To ensure homogeneous dispersion of the genomic DNA: (a) allow the DNA to stand at 4°C for several hours after dilution and addition of 10 x restriction enzyme buffer, (b) gently stir the DNA solution from time to time, (c) after adding restriction enzyme, gently stir the solution for 2-3 minutes at 4°C before warming up to 37°C, (d) after digestion for 30 minutes, add a second aliquot of restriction enzyme and stir as described above)
Add 0.1 volume (10µl) 3 M sodium acetate, mix well. Add 2.5 volumes ice-cold 100% ethanol. Precipitate at -80°C for 1 hour. Spin in microfuge at high speed for 20 minutes. Discard supernatant, air-dry for 5 minutes. Resuspend in 15 µl 10mM Tris pH 7.5.
(Residual ethanol can cause the DNA sample to 'crawl' out of the well when loaded onto the gel. Heating the solution of redissolved DNA to 65-70°C in an open tube for 10 minutes can usually drive off most of the ethanol.)
Add 3 µl 6 x DNA-loading buffer (total volume 18 µl).
Pour a 0.8% agarose gel.
(Mix 1.6 grams ultra pure agarose in 200 ml 1X TAE (0.8%). Add 2 µl 1% ethidium bromide solution. Pour into long (20 cm) gel rig)
Prepare DNA size markers.
(2µl lambda HindIII (1µg) + 1µl 100 bp ladder (1µg), 9µl H2O, 3µl 6x DNA loading buffer)
Optional: Prepare positive control.
(Mix 10µg digested genomic DNA with 1-10 pg plasmid containing the cDNA for which will be probed. Load into lane far outside from samples to avoid contamination or interference with sample hybridization)
Submerge gel in 2000 ml 1X TAE with 0.5 µg/ml ethidium bromide.
(i.e. 40 ml of 50 x TAE, 100 µl of 1% ethidium bromide stock solution, dH2O ad 2 liter)
Load samples, markers and controls.
(If DNA does not sink to the bottom of the well, make sure that all the ethanol was removed. Load samples very slowl. After loading, allow the gel to stand for a few minutes so that the DNA can diffuse evenly troughout the wells.)
Electrophorese at 40 Volts, 35 mAmps for 6-16 hours.
Photograph with fluorescent ruler 1/8th sec with ruler and an additional 1 sec on gel alone. Cut off the buttom left-hand corner of the gel helping orientation.
For partial fragmentation of large DNA fragments (>10kb) prior to the transfer, leave on UV light box for 10 minutes.
(Alternatively, partial fragmentation can be achieved by acid depurination: Invert gel, place into 0.25M HCl for 10 min, rinse briefly in dH2O.)
Just prior to the transfer, float the Nylon membrane (S&S Maximum Strength Nytran) on top of distilled water to wet thoroughly. Let stand in water or transfer buffer until use. Cut off lower left-hand corner for orientation.
Denaturation: Soak gel in at least two gel volumes (500 ml) of 1.5 M NaCl / 0.5N NaOH for 2 x 15 minutes.
If gel floats to the surface of the liquid, weigh it down with several pasteur pipets.
Set up transfer in large electrophoresis tray: Fill transfer buffer (10 x SSC) into the two side-trays. Place the following items (gel-sized, saturated with transfer buffer) on the middle support. All air-bubbles should be carefully smoothed-out with a glass pipet or gloved fingertip in each layer seperately.
Wick (3mm paper); this should be the same width as your gel and long enough to drape well into the transfer solution.
3 pieces of Whatman 3MM paper
Gel (upside down)
Nylon membrane (Always handle with clean gloves and blunt-ended forceps. Do not adjust the Nylon membrane once it is placed on the gel)
3 pieces of Whatman 3MM paper
dry 5 cm stack of paper towels
glass plate with weight (ca. 500 g)
Surround gel with plastic wrap or parafilm to prevent the paper towels to come in contact with the wet paper below the gel. Prevent evaporation of the transfer solution by sealing with plastic wrap or parafilm on either end of the tray. Let transfer overnight.
Next day, take off blotting material and mark the position of the wells with very-soft-lead pencil. Soak the gel in 5 x SSC for 5 minutes to remove bits of gel or particles from membrane.
Place wet membrane on a paper towel of equal size. Immobilize the DNA by UV-crosslinking (100 mJ/cm2, push "optimal cross-linking"). Dry membrane thoroughly at room temperature before placing in prehybridization solution.
Place membrane in hybridization bottle with 20 ml (min. 100 µl/cm2) of prehybridization buffer (use nylon mesh as a spacer before rolling). Incubate in hybridization oven at 42°C for 1-2 hours. Alternatively, without formamide, incubate at 68°C. Some workers leave out Denhardt's in hybridization and /or prehybridization solution.
Prehybridization Buffer (20 ml)
|deionized formamide||50%||10 ml|
|20 x SSPE stock solution||6 x||6 ml|
|50 x Denhardt's reagent||5 x||2 ml|
|20% SDS stock solution||0.5%||500 µl|
|Salmon Testes DNA (Sigma D-9156), |
10.4 mg/ml, boil 5 min. before use
|100 µg/ml||200 µl|
|optional: dextrane sulfate||10%||2 g|
|dH2O ad 20 ml||1.3 ml|
Isolate 1-10 µg target cDNA from vector with restriction digestion, agarose gel electrophoresis, gel extraction of appropriate band. Resuspend in 10 mM Tris.
Label 25-50 ng cDNA-probe with 32P using the Prime It® II Random Primer Labeling Kit (Stratagene #300385).
Add to the bottom of a microfuge tube:
Boil reaction tube in water bath for 5 minutes, then centrifuge briefly at room temperature to collect condensed liquid from cap of tube.
Add the following reagents to the reaction tube:
Mix the reaction components thoroughly with pipet tip.
Add denatured probe to prehybridization solution.
(If Denhardt's was in the prehybridization solution and should not be in the hybridization solution, prepare fresh prehybridization solution without Denhardt's for hybridization.)
Incubate overnight (12-24 hours) at 42°C (or at 68°C without formamide) in hybridization oven.
The hybridization solution can be reused.
(Pour into a 50 ml plastic disposable centrifuge tube. The probe is good for a couple of weeks, and must be boiled 5 min and chilled on ice before reuse.)
Soak membrane twice for at least 15 minutes each with 100 ml of 7 x SSPE / 0.1-0.5% SDS at room temperature. Perform this step in hybridization oven rotating at very low speed.
(for 500 ml: 175 ml 20 x SSPE, 2,5 - 12,5 ml 20% SDS, 322.5 - 312.5 ml dH2O)
Then soak the membrane twice for at least 15 minutes each in 100 ml of 1 x SSPE / 0.5-1% SDS at 37°C. Perform this step in hybridization oven rotating at very low speed.
(for 500 ml: 25 ml 20 x SSPE, 12.5 - 25 ml 20% SDS, 462.5 - 450 ml dH2O)
Finally soak for 1 hour in 100 ml of 0.1 x SSPE / 1% SDS at 68°C.
(for 500 ml: 2.5 ml 20 x SSPE, 25 ml 20% SDS, 472.5 ml dH2O)
For detection of poorly matched hybrids use a lower temperature in the last wash. Check radioactivity with Geiger-counter. Do not expect a signal from single copy genes.
After washing, blot the membrane with filter paper (Whatman 3MM or S&S GB003) to remove most of the excess moisture. Wrap moist blots in plastic wrap prior to autoradiography. Expose to X-ray film with an intensifying screen at -80°C for 1 to 6 days.
Do not allow blot to dry at any time prior to removing the probe, as drying will cause the probe to bind irreversibly. To remove probe from blot for reuse. incubate one of the following solutions:
Wash blots in 50% formamide, 6 x SSPE for 30 minutes at 65°C.
(for 50 ml: 25 ml deionized formamide, 15 ml 20 x SSPE, 10 ml dH2O)
Rinse in 2 x SSPE, remove excess liquid by dabbing the blot on 3MM paper. Wrap in Saran wrap and store. Expose overnight to check for the absence of radioactivity. After stripping probe, start again with prehybridizing step.