High molecular weight DNA was obtained by phenol extraction, using a method adapted from that of Kunkel et a l, (1985). Approximately 0.5 g tissue was
homogenised in 4 ml TE (10 mM Tris-HCl pH 8.0, 1 mM EDTA pH 8.0) and protein digestion was carried out at 55°C overnight by adding 0.5 ml 0.5 M EDTA pH 8.0, 50 pi 10% SDS and 100 pi 10 mg/ml proteinase K. The digests were extracted three times
with an equal volume of equilibrated phenol, to remove any remaining protein, and then twice with chloroform, to remove phenol. At each step the samples were mixed gently to avoid shearing the DNA, then centrifuged at 10,000 rpm for 10 minutes at room temperature to separate the aqueous (upper) phase from the organic (lower) phase. The aqueous phase was removed each time with a phenol hook, taking care to leave behind the proteinaceous interface. DNA was precipitated at room temperature from the final aqueous phase by the addition of two volumes of 100% ethanol, and the stringy precipitate was spooled out using a phenol hook, washed in 70% ethanol, and resuspended in 2 ml TE buffer. DNA concentrations and purity were determined spectrophotometrically by measuring the optical density (OD) at 260 and 280 nm. The OD at 260 nm was used to calculate the DNA concentration (one OD unit = 50 pg/ml), and the 260/280 ratio used to determine the purity (pure DNA gives a ratio of 1.8). DNA samples were subsequently diluted to 200 pg/ml in TE buffer and stored at 4°C.
2.7.2 Restriction enzyme digestion of DNA
Restriction digests of DNA samples were carried out in accordance with the conditions suggested by the manufacturers (Promega, Gibco BRL or New England Biolabs). Analysis of genomic DNA was typically performed with 5 pg DNA and 1.25 mM spermidine in a total volume of 80 pi. Up to 20 units of enzyme were added at the beginning of the 6-8 hour incubation at the recommended temperature. A further 10 units were added after about 3 hours, to boost digestion. Digestion of DNA samples for other reasons, for example, digestion of plasmid DNA, was carried out in essentially the same way, but usually in a smaller volume, with a smaller excess of enzyme and for less time (~2 hours), depending on the mass of DNA being digested.
2.7.3 Electrophoresis of genomic DNA digests
Size fractionation of the products of genomic DNA digests, for Southern blotting, was performed by agarose gel electrophoresis, essentially as described in section 2.5.
Adequate separation of the high molecular weight (1-15 kb) products required the use of large (20 x 20 cm) 0.8% agarose gels, and electrophoresis was carried out at 55 volts (2.75 V/cm) for 16-20 hours. Gels were photographed on a UV light-box, with a ruler alongside to allow the later calculation of fragment sizes on the Southern blot.
2.7.4 Southern blotting with 20 x SSC
This procedure was carried out essentially as described by Southern (1975), and results in the transfer of size fractionated DNA fragments from an agarose gel to a nylon membrane. Gels to be blotted were placed, after photography, in denaturing solution (0.15 M NaOH, 0.015 M sodium citrate) and shaken for approximately 1 hour at room temperature, depending on the gel size and thickness. This denatures the double stranded DNA making it accessible for hybridisation. After dénaturation, the gels were transferred into neutralising solution (3.0 M NaCl, 0.5 M Tris-base, pH 5.5), for an equal time period, again with shaking.
The gel was inverted and placed on a triple layer of Whatmann 3MM paper set up on a support (often a gel former) in a tray of 20 x SSC (3 M NaCl, 0.3 M sodium citrate, pH 7.0). The ends of the Whatmann paper were allowed to dip into the SSC, to act as a wick. Strips of Nescofilm or Saran Wrap were placed along the edges of the gel to prevent short-circuiting of the SSC directly into the paper towels, and either Hybond N (Amersham) or Electran nylon membrane (BDH) cut to the size of the gel was prewetted in 3 X SSC and positioned on top of the gel. Three sheets of Whatmann 3MM paper,
soaked in 3 x SSC, were layered on top of the nylon membrane. Care was taken to exclude air bubbles and ensure good contact between layers, by gently rolling the stack after each layer was added. A stack of paper towels and a weight were then placed on top of the Whatmann paper, and the blotting procedure was allowed to take place over
14-16 hours. Capillary action results in movement of SSC from the tray into the paper towels taking with it the DNA from the gel, and under these high salt conditions the DNA binds to the membrane. On completion of blotting, the blot was dismantled and the positions of the wells were marked on the membrane before briefly rinsing in 3 x SSC. The DNA was permanently fixed to the membrane by ultraviolet (UV) irradiation in a Stratagene UV Crosslinker, following manufacturers guidelines, and the blots were stored dry.
2.7.5 Oligonucleotide labelling of double stranded DNA probes
DNA probes were labelled to a high specific activity using the random hexanucleotide priming method (Feinberg and Vogelstein, 1984). Denatured double stranded DNA fragments were annealed to random hexanucleotide primers, then the addition of the Klenow fragment of DNA polymerase I and nucleotides dATP, dOTP, dTTP with
radiolabelled dCTP resulted in the synthesis of a new strand incorporating the labelled nucleotide.
DNA (50-150 ng) in a volume of 35 pi was denatured by heating to 95°C for 7 minutes, then quenched on ice. The labelling reaction was set up by adding 10 pi 5x Oligo Labelling Buffer, 1 pi Klenow (5 Units; Bioline) and 4 pi (40pCi) [a-^^P]-dCTP (3000 Ci/mmol, 10 mCi/ml; ICN) to the denatured DNA, to give a final volume of 50pl. This mixture was incubated at 37°C for 1-3 hours, then unincorporated [a-^^P]-dCTP was removed by purification through a Sephadex G50 spin column (see section 2.7.6).
Oligo Labelling Buffer is A : B : C in the ratio 1 : 2.5 : 1.5 (stored at -20°C), where:- Solution O = 1.25 M Tris HCl, pH 8.0, 0.125 M MgCl2
Solution A = 1 ml Solution O plus: 5 pi each of 0.1 M dATP, dGTP and dTTP, and 18 pi p-mercaptoethanol
Solution B = 2 mM A^-2-hydroxyethylpiperazine-A^’-2-ethanesulfonic acid [HEPES]
pH 6.6
Solution C = Hexadeoxyribonucleotides (Gibco BRL) in TE buffer at 90 OD units/ml
2.7.6 Separation of unincorporated nucleotides from the labelled probe
Unincorporated nucleotides were removed from the labelled probe by fractionation through a Sephadex G-50 spin column. The labelled probe passes quickly through the column, while the unincorporated nucleotides move more slowly through the column matrix, enabling the probe to be separated. Spin columns were prepared in the barrels of 1 ml disposable syringes, plugged with polymer wool and filled with a slurry of 8% (w/v) Sephadex G-50 (Sigma) in TE. The column was centrifuged at 1750 rpm for 4 minutes to settle the Sephadex, then equilibrated by spinning through 200 pi of ddH20. The labelling reaction was made up to 200 pi with ddH20 then spun through the column. The filtrate (the labelled probe) was carefully collected and the activity assessed by counting a 2 pi aliquot with a Bioscan GC2000 radiation counter. The probe needs to be labelled with high enough specific activity to give approximately 0.5
million counts per minute (cpm)/ml hybridisation mix. Labelled probes were either used immediately or stored in shielded boxes at -20°C for up to 1 week.
2.7.7 Prehybridisation of Southern blots
Southern blots, interleaved with muslin sheets to separate overlapping membranes, were wetted in 6 X SSC then rolled up and placed into hybridisation roller bottles.
Membranes were prehybridised in 20-40 ml prehybridisation solution (6 x SSC, 0.5% SDS, 5 mM EDTA pH 8.0, 5 x Denhardts solution [50 x Denhardts solution is 1% w/v bovine serum albumin (BSA), 1% w/v ficoll and 1% w/v polyvinylpyrrolidone (PVP)], 0.1 mg/ml denatured salmon sperm DNA) at 65°C, with rotation, for 4-16 hours.
2.7.8 Hybridisation of Southern blots
Following prehybridisation, the prehybridisation solution was discarded and replaced with 8-15 ml hybridisation solution (prehybridisation solution with the addition of 10% w/v dextran sulphate). DNA probes were denatured by heating to 95°C for 10 minutes, quenched on ice, then added to the hybridisation mix. Hybridisation was carried out for 12-36 hours, at 65°C.
2.7.9 Removal of unbound probe
Following hybridisation, the hybridisation solution containing unbound probe was carefully discarded and the bottle and membranes rinsed with 3 x SSC/0.1 % SDS. The membranes were washed in 100-200 ml 3 x SSC/0.1 % SDS at 65°C for 30 minutes. Following this wash, the membranes were removed from the bottle, and the activity monitored with a Geiger counter. If the activity was > 1 0 counts per second (cps), the membranes were washed more stringently, by decreasing the SSC concentration (the SDS concentration was kept at 0.1%). The usual series of washes used was 3 x, 2 x, 1
X, 0.5 X, 0.25 X, and finally 0.1 x SSC, with the membranes monitored after each wash
until the activity had been reduced to 5-10 cps. The membranes were then removed from the wash solutions, sealed between 2 sheets of Saran wrap and placed into an autoradiography cassette with intensifying screens. The membranes were
autoradiographed using either X-omat hyperfast film (Kodak) or blue sensitive X-ray film (Genetic Research Instrumentation), at -70°C, with an exposure time of between 16
hours and 2 weeks, depending on the intensity of the signal. Autoradiographs were developed using an RGII X-ray film processor (Fuji).
2.7.10 Stripping Southern blots for reprobing
Membranes were stripped to remove all the bound probe, allowing them to be successfully reused four or five times. Stripping was performed by washing the
membrane in 0.4 M NaOH at 45°C for 30 minutes, then in 0.1 x SSC/0.1% SDS/0.2 M Tris pH 7.5 for 15 minutes at 45°C. Before reprobing, the membranes were
autoradiographed overnight to check the stripping process was complete.