There are many different protocols for Southern blotting. This approach uses vacuum transfer of DNA to a nylon membrane, DIG-labelling of the probe and detection by chromogenic substrate or chemiluminescence. The buffers for DNA transfer are as described by Sambrook et al., transfer is carried out by following the instructions for the vacuum pump. The protocols for labelling and detection are as described by the manufacturer’s instructions (Roche Life science in this case).
Consumables and Solutions
Labelling the Probe
If a blot fails completely, it will almost always be because of a bad probe. For best results: label around 1 µg DNA. Two options for DIG-labelling are described here:
- Random primed DNA labelling: To label a whole cosmid or a fragment of DNA (e.g. fragment excised from a plasmid or a PCR product).You will need a very concentrated DNA sample for this, as each reaction will only label up to 15 µl DNA, and you want approximately 1 µg DNA.
- PCR labelling: Labels during PCR and therefore generates a lot of labelled DNA from small amount of template. This is a more expensive method but, provided PCR has already been optimised, is faster and more likely to generate the required amount of probe.
1. Random Primed DNA Labelling
In this method, DNA is made single stranded and then hexanucleotides act as random primers for second strand synthesis using Klenow enzyme. The DIG DNA labelling mix contains dATP. dCTP, dGTP and DIG-labelled dUTP, so DIG is incorporated into the second strand every 20-25 nucleotides as it is synthesised.
- Make volume of DNA sample to 15 µl with water (10 ng – 3µg).
- Boil at 95°C for 10 min to denature DNA (can use a PCR machine for best results), then chill quickly in an ice water bath for around 10 min.
- Set up the following reaction:
- 15 µl denatured DNA
- 2 µl Hexanucleotide mix 10x
- 2 µl DIG DNA labelling mix
- 1 µl Klenow enzyme
- Mix and incubate at 37°C for 1h-20h (for best results, leave overnight)
- Stop the reaction by adding 2 µl 0.2M EDTA (pH 8.0).
- Store at -20°C until required.
2. PCR Labelling
The PCR DIG labelling mix contains dATP, dCTP, dGTP and DIG-labelled dUTP, so as the DNA is amplified in a standard PCR reaction DIG is incorporated into every product.
- Set up PCR reaction as usual but instead of using standard dNTPs use PCR DIG labelling mix. Mix is 10x concentrated so use 5 µl per 50 µl reaction.
- Labelled PCR products can be used directly as probe. However, if there are any non-specific products, the correct size product should be excised from an agarose gel and eluted before use.
- You can combine the products of several reactions to use as your probe.
- Store at -20°C until required.
Note: You will need to prepare a probe for the DNA ladder as well in order to have a size reference on the blot. You can prepare your own Lambda DNA/HindIII Ladder and label with DIG by Random Primed DNA Labelling.
- Digest DNA samples as required and separate on a 0.6-0.8% agarose gel for 4-6 h or overnight.
- Try to add the same amount of DNA to each lane.
- If using a positive control, consider the copy number of your target fragment. For example, if you are using digested plasmid as your positive control and digested genomic DNA in your experimental lanes, you will need to add significantly less plasmid DNA than genomic DNA. As a rule of thumb, dilute your plasmid DNA so that you can just barely visualise it on your gel, if you add too much positive control the probe and antibody will be sequestered by this DNA and will give too bright a signal compared to experimental lanes.
- Similarly, dilute your ladder considerably to avoid overdetection of blotted DNA ladder.
- Consider which enzymes to digest with carefully. It will be difficult to differentiate the sizes of large fragments on your blot, so if possible, aim for fragment sizes of <7 kb.
- Be sure to run your DNA far enough through the gel for best resolution of target fragments.
- You can either add ethidium bromide to the gel or stain after electrophoresis. Either way, take a photo as usual (before the blotting) to refer to later.
Transfer to Membrane
- Measure the area of the gel that you want to be transferred to the membrane. You can trim the gel so that it has a border around this area of approximately 2 cm. There must be an overlap otherwise vacuum will not form properly.
- Prepare the following before proceeding:
- Cut three sheets of Whatman paper approx. same size as the gel.
- Cut plastic sheeting so that it covers the entire platform of the transfer tray, and cut out a transfer window from the middle of the plastic, exactly the size determined in Step 1.
- Cut membrane so that it overlaps the transfer window by approximately 1 cm all the way round. NB Handle the membrane as little as possible, only at the edges, and only gloved hands.
- Use a pencil to label the back of the membrane, in the top right corner. NB It is very important to keep track of witch side of the membrane is the front.
- Prepare the transfer apparatus as follows:
- Clean the base sheet and place on tray, smooth side up.
- Soak the three sheets of Whatman paper in 2x SSC and place on the centre of base sheet.
- Carefully place the membrane on top (with writing facing down).
- Place plastic sheet over, with transfer window central to the membrane. There should be an overlap of approx. 1 cm. Clip lid of transfer tray into place.
- To check the seal:
- Clean glass sheet with 2x SSC and lay over transfer window.
- Connect transfer tray to pump and switch it on.
- Vacuum pressure should stabilise at around 50mbar. The dial can be used to adjust the pump strength, but if pressure continues to fall it means the seal is faulty. If it happens, it is probably because the plastic sheeting is not covering the platform sufficiently. Take the assembly apart and try a bigger piece of plastic if necessary.
- Switch off pump and remove plate when satisfied that the seal is sufficient to create a vacuum.
- Carefully slide your gel onto the assembly, ensuring that the transfer window aligns to the right section of the gel. Try to avoid air bubbles, rips and tears, but you can remove any air bubbles by smoothing out with a gloved finger.
- Pipette 50 ml depurination buffer onto gel and switch on pump for around 20 min.
- Switch off pump and remove depurination buffer by tilting assembly and then using pipette to remove buffer from corner of tray.
- Pipette 50 ml denaturation buffer onto gel and switch on pump for 20 min.
- Switch off pump and remove buffer as described in Step 7.
- Pipette 50 ml neutralisation buffer onto gel and switch on pump for 20 min.
- Switch off pump and remove buffer as described in Step 7.
- Pipette 50 ml 20x SSC onto gel and switch on pump for around 1 hour to allow DNA transfer to occur.
- Disassemble blot and carefully wrap membrane in clingfilm. Expose to UV for 2 min to crosslink.
- Remove from clingfilm and wash membrane in 2x SSC for 2 min.
- Wrap membrane in a piece of mesh and insert into Hybaid tube. NB If the mesh overlaps with the top of the tube it will break the seal and allow buffer to leak.
- Add 20 ml preheated (~65°C) SHB solution and prehyb in rotating Hybaid oven at 65°C for 1 hour.
- Defrost probe and denature at 100°C for 15 min, then put on ice for 10 min. If reusing old probe, you can denature probe in a Falcon tube, within a beaker of water over a hot plate.
- Decant off old SHB (prehyb) and replace with 20 ml fresh SHB solution plus denatured probe.
- Hybridise overnight at 65°C.
Note: SHB + probe can be stored at -20°C and reused several times when freshly denatured at ~65°C for 10 min before use.
- Decant SHB and probe from Hybaid tube. The probe can be frozen and reused (store in a Falcon tube).
- Add ~30 ml Stringency Buffer A to Hybaid tube and rinse.
- Wash membrane in Hybaid tube for 15 min with ~150 ml Stringency Buffer A, in Hybaid oven at 65°C.
- Repeat Buffer A wash.
- Add ~30 ml Stringency Buffer B to Hybaid tube and rinse.
- Wash membrane in Hybaid tube for 15 min with ~150 ml Stringency Buffer B, in Hybaid oven at 65°C.
- Repeat Buffer B wash.
- Remove membrane and mesh from Hybaid tube and rinse in Washing buffer for 20s – 30s (no longer).
- Wash for 30 min in 100 ml freshly prepared 1% blocking solution in a clean tray, with gentle rocking or shaking. (This blocks unspecific binding sites on membrane)
- Spin anti-DIG-AP solution for 10 min (reduces speckling on blot).
- Pour off blocking solution and replace with 30 ml antibody solution. Antibody binds to the DIG-labelled probe. Incubate for 30 min with gentle rocking or shaking.
- Decant, and wash in ~100 ml Washing buffer for 15 min to remove unbound antibody.
- Repeat wash.
- Decant washing buffer and equilibrate for 2 min in 20 ml Detection buffer.
- Incubate membrane in 10 ml freshly prepared Colour-substrate solution: 200µl of NBT/BCIP in 10 ml of Detection buffer, kept away from light!), in an appropriate container in the dark. Do not shake during colour development.
- Colour usually develops within a few minutes and the reaction is usually complete after 16 h. The membrane can be exposed to light for short time periods to monitor colour development.
- Stop the reaction when desired spot or band intensities are achieved, by washing the membrane for 5 min with 50 ml of sterile double distilled water or TE-buffer.