John Wiley & Sons - 2004 - Analysis of Genes and Genomes

Mix and pour into block former. Allow to cool

Incubate with hydrolytic enzymes

Digestion of cells

DNA trapped

Agarose blocks of DNA loaded into agarose gel slots

Figure 2.20. Preparation of high-molecular-weight DNA for analysis by PFGE. Intact cells are mixed with molten, but cool, agarose and poured into a block former. Once set, the agarose blocks can be treated with hydrolytic enzymes to break open the cell walls and release the chromosomal DNA. If required, the DNA can be digested with restriction enzymes whilst also in the agarose block. The treated block is then loaded into the well of an agarose gel before being subjected to PFGE

that the sequence of the DNA in each band on the gel is different to other bands – even bands that are identical in length may have a very different DNA sequence. It is, of course, the DNA sequence itself that plays a vital role in the function of a molecule. In the mid-1970s methods were developed to distinguish

bands on gels that contain a particular DNA sequence. These methods rely on the hybridization of nucleic acid sequences in order to detect the presence of complementary sequences. The original method of blotting was developed by Ed Southern in 1975 for detecting DNA fragments in an agarose gel that were complementary to a given nucleic acid sequence (Southern, 1975).

2.6.1Southern Blotting

In the procedure, referred to as Southern blotting, the DNA fragments separated on an agarose gel are transferred and immobilized onto a membrane. After the gel has been run, the DNA fragments are denatured (i.e. the strands are separated) using alkali. The single-stranded nature of the DNA on the membrane is important to allow complementary DNA sequences to be able to bind to the DNA fragments attached to the membrane. After the DNA in the gel has been denatured, the DNA fragments are transferred to a membrane. The transfer process can be mediated either electrophoretically or through capillary attraction by placing the gel and membrane in contact with each other and allowing buffer to flow through the gel onto the membrane – the DNA fragments move with the buffer and become trapped on the membrane. Initially, nitrocellulose membranes were used, but these were fragile and easily broken. Nylon membranes are commonly used today. After transfer, the DNA

fragments need to be fixed to the membrane so that they cannot detach. A number of methods of fixing are available including baking at 80 ◦C and ultraviolet cross-linking. UV cross-linking is based on the formation of crosslinks between a small fraction of the T residues in the DNA and the positively charged and amino groups on the surface of the nylon membrane. Following fixation, the membrane is placed in a solution of labelled (often radioactive) single-stranded nucleic acid – either single-stranded DNA or RNA. The labelled nucleic acid (or probe) is allowed to hybridize to its complementary partner sequence on the membrane. The interaction between the single-stranded probe and its complementary sequence will result in the binding of the probe to the membrane through non-covalent hydrogen bonding that normally holds the DNA double helix together. The membrane is then washed extensively to remove non-specifically bound probe, and specific interactions are detected by exposing the membrane to X-ray film (Figure 2.21).

Southern blotting is used to detect DNA sequences that are either identical or similar to the sequence of the probe. The hybridization of the probe to the DNA sequences trapped on the membrane is the critical component of success of these experiments. As we have seen previously, single-stranded DNA will bind with high affinity to its complementary partner sequence. It will also

DNA separated through agarose gel

Blot DNA onto membrane

Add single-stranded labelled DNA

Expose membrane to X-ray film

Band on film shows position of complementary DNA sequences

Figure 2.21. Southern blotting. DNA fragments separated on an agarose gel are transferred onto a membrane. The DNA is made single stranded before the addition of a labelled probe. The binding of the probe to the membrane – through base pair hydrogen bonding – can be detected by exposing the membrane to X-ray film. The radio-labelled probe will form a band on the film at a position corresponding to the complementary sequence on the membrane

bind to similar, but non-identical, DNA sequences with a reduced affinity. This differential binding affinity to different DNA sequences can be used to identify DNA molecules that are not identical but are merely related to the probe. This sort of analysis is achieved by altering the temperature (or salt concentration) at which the probe is washed off the filters after it has bound (Figure 2.22). Washing the membrane at high temperature (high stringency) will result in thermal disruption of all but the most tightly bound sequences. Consequently, the bands that show up on the X-ray will film will be either identical or highly related to the probe. Washing the membrane to lower temperatures will reduced the overall level of stringency, and sequences that are less related to the probe

Add single-stranded labelled DNA

Figure 2.22. Washing Southern blot membranes at different temperatures results in different stringencies. Washing the membrane at high temperature will remove all but the most tightly bound DNA molecules – those most similar to the sequence of the probe. Lower-temperature washes can reveal sequences on the membrane that are similar, but not identical, to the sequence of the probe. The actual temperature at which washes are performed will depend on the length of the probe and the likelihood of it finding an exactly matching sequence on the membrane

will give positive signals on the X-ray film. This approach is immensely useful when you do not precisely know the sequence of the gene you wish to identify. This situation often arises when knowledge of the protein sequence is known, but the DNA sequence encoding that protein is unknown. We shall discuss this issue further in Chapter 6.

2.6.2The Compass Points of Blotting

Since Southern’s first description of blotting and hybridization, a number of variants have been described. These all involve the transfer of material from a

Proteins separated through polyacrylamide gel

Blot proteins onto membrane

Block non-specific protein binding sites and add primary antibody

Antibody binds to antigen

Wash away excess primary and add labelled secondary antibody

Antibody sandwich

Detect label

Band shows position of antigen

Figure 2.23. Western blotting using antibodies to detect specific proteins. Proteins are separated through a polyacrylamide gel, usually containing SDS to maintain them in a denatured state, prior to blotting onto a nitrocellulose membrane. Non-specific protein binding sites are blocked on the membrane – using solubilized milk powder – before the primary antibody is added. The primary antibody will specifically bind to the antigen to which it was raised. A labelled secondary antibody is then added to detect the location of the primary antibody. The antibody sandwich results in an amplification of the signal. The secondary antibody is often labelled with an enzyme whose activity, in the presence of appropriate substrates, results in either a colour change on the membrane or the emission of light that can be detected using X-ray film

the cell wall will result in the nuclear material spilling out from the broken cells. This material can be harvested, but the DNA will be contaminated with both RNA and with proteins. Methods are therefore required to remove these contaminants to yield a purified DNA fraction.

Total genomic DNA isolation is often carried out according to the following procedure.

•The host cell wall is lysed. The method used for the lysis procedure depends upon the nature of the host cell itself. For instance, bacterial cells are often treated with the enzyme lysozyme to weaken their cell wall before being lysed with detergents. Yeast cells, on the other hand, are treated with zymolase to disrupt the integrity of their cell wall before lysis proceeds – often by grinding the cells using glass beads to break them open.

•The cellular debris will be removed by centrifugation.

•The remaining soluble material will then be vortexed in the presence of phenol to remove proteins by denaturing then. Chloroform is often used in conjunction with phenol (as a phenol/chloroform solution) since it is also a protein denaturant, but it also stabilizes the rather unstable boundary between the aqueous phase and a pure phenol layer.

•The nucleic acid remaining in the aqueous layer can then be precipitated using ethanol. Upon addition of ethanol the DNA will form a stringy white precipitate that can be collected by a centrifugation.

•RNA can be removed from the preparation by treatment with RNase.

For many applications, the DNA must be purified further. Isopycnic centrifugation can be used to separate DNA from contaminants, and also to isolate individual types of DNA molecule. The DNA sample is mixed with a substance (caesium chloride, CsCl) that is capable of forming a self-generating uniform density gradient during the centrifugation process. DNA molecules will sediment only to the position in the centrifuge tube at which the gradient density is equal to its own density, and they will remain there to form a band. After the DNA band has formed, it is removed from the centrifuge tube and the CsCl removed by precipitating the DNA with alcohol. A major advantage of this technique is the ability to separate different species of DNA molecules (Figure 2.24). The inclusion of ethidium bromide in the centrifugation mix results in its binding to DNA. The topological constraints of supercoiled DNA mean that closed-circular DNA molecules (e.g. plasmids) are able to bind only limited amounts of ethidium bromide. Nicked DNA circles, and linear DNA fragments, are able to bind much more ethidium bromide and are extensively unwound in its presence (Figure 2.12). This results in nicked and linear DNA being less dense than their supercoiled equivalent, and consequently being separated in a different location on a CsCl/EtBr gradient. The main drawback of isopycnic centrifugation is the long hours of centrifugation required to form

Bacterial cell

SDS + sodium hydroxide

Neutralise with potassium acetate

Genomic DNA and proteins precipitated

Remove precipitate by centrifugation

Plasmid DNA in solution

Bind to gel matrix

Elute impurities

Elute DNA

Pure plasmid DNA

Figure 2.25. Plasmid DNA purification. Bacterial cells harbouring plasmid DNA are broken open using SDS. The presence of sodium hydroxide ensures the denaturation of genomic DNA. Upon neutralization, the genomic DNA aggregates and forms a precipitate together with the SDS–protein complexes. These can be removed by centrifugation. The plasmid DNA is then bound to a matrix to which contaminants are unable to adhere. The pure plasmid DNA can then be eluted from the matrix