- 1 DNA Sequencing by primed synthesis methods
- 2 Principle of the ‘plus and minus’ method
- 3 Additional methods useful in conjunction with plus and minus method
- 4 Procedure
DNA Sequencing by primed synthesis methods
DNA sequencing method is used for the determination of two particular nucleotide sequences in bacteriophage DNA using DNA polymerase primed by synthetic oligonucleotides. The target oligonucleotide is hybridized to a specific complementary region on the single strand DNA. Deoxynucleotides is added sequentially by DNA polymerase to the 3‘-OH end of the primer. Using [32P]-labelled deoxyribonucleoside 5‘-triphosphates a radioactive complimentary copy of a defined region of the template is obtained.
Next stage of development on this method came after two years. Analysis of the complementary DNA is done by fractionating the DNA fragments by electrophoresis on high resolution by polyacrylamide gels. It constitutes a relatively rapid and simple method for sequence analysis and illustrates the principle on which the modern chain-termination procedure is based.
Principle of the ‘plus and minus’ method
A DNA primer (commonly a restriction fragment or synthetic oligonucleotide) is hybridized to the single-stranded DNA template and the primer is extended to a limited degree with DNA polymerase I in the presence of all four deoxynucleoside 5‘- triphosphates one of which is labelled. Samples of the reaction mixtures are taken at different time periods to find the degree of extension of primers. The reaction is terminated by addition of EDTA and the various samples are then combined. DNA polymerase is removed by phenol extraction. Extended polynucleotide chain, still hybridized to the template, is separated from the excess deoxynucleoside triphosphates by gel filtration on sephadex or agarose. The product is a mixture of partially elongated fragments with variable chain length. Ideally the product is a mixture of polynucleotides in which all possible chain lengths of the complementary strand are present corresponding to an elongation of the primer from 0 to 200 nucleotides.
The ‘minus’ reaction
Due to the absence of one of the four deoxynucleoside triphosphates in this reaction, DNA polymerase would accurately catalyze chain extension up to the point where the missing nucleotide should have been incorporated. The ‘minus’ reaction utilizes the same principle. The random mixture of oligonucleotides, still annealed to the DNA template is re-incubated with DNA polymerase in the presence of only three deoxynucleoside triphosphates. Synthesis then proceeds as far as the missing triphosphate on each chain. Four separate reactions are set up, each with one of the triphosphates missing. After incubation the reaction mixtures are denatured to separate the nascent strands from the template and the four samples simultaneously analyzed by electrophoresis on a high resolution polyacrylamide gel in the presence of 8M urea and the separated oligonucleotides visualized by autoradiography.
On the acrylamide gel mobility is essentially proportional to size so that the various oligonucleotide products, all of which should have a common S-end will be arranged in a ladder according to size. Each oligonucleotide should be resolved from its neighbor, which contains one more residue, by a distinct space. As would be expected the resolution falls off with increasing chain length and it is this factor which usually determines how far the autoradiograph may be read. The separation between two fragments of say 150 and 151 nucleotides will be much less than that between two fragments of 20 and 21 nucleotides. The autoradiograph of the -A channel will consist of a set of bands, each of which corresponds to an extension product up to, but not including, the next A residue in the sequence. Thus the positions of the A residues are located. In a similar way the positions of the other residues are located from the sequencing channels and, in principle, the sequence of the DNA is read off from the autoradiograph. Usually, however, this system alone is not sufficient to establish the sequence and a second line of attack, the ‘plus’ system is used to confirm and complement the data from the ‘minus’ system.
The ‘plus’ system
In the presence of a single deoxynucleoside triphosphate, DNA polymerase from phage T4 infected E. coli (T4 polymerase) will degrade double-stranded DNA from its 3‘-ends but this 3’-exonuclease activity will stop at residues corresponding to the single deoxynucleoside triphosphate present in the reaction mixture. Since T4 polymerase lacks the 5’-exonuclease activity found in E. coli polymerase incubation with this enzyme serves to trim back each elongated product to the residue corresponding to the added deoxynucleotide. The polymerizing activity of the enzyme catalyses the turnover of this residue but effectively halts the progress of the 3’-exonuclease activity. In the ‘plus’ reaction this method is applied to four further samples of the primertemplate complex isolated above. Samples are incubated with T4 polymerase and a single triphosphate and, after denaturation, the products are analysed directly by gel electrophoresis. Thus, in the +A system only dATP is added and all the chains will consequently terminate with a terminal A residue. The bands observed on the gel will therefore correspond to a set of fragments representing all the extension products which terminate with A. The products from the ‘plus’ reaction will be one residue longer than the corresponding band in the ‘minus’ A reaction.
The polyacrylamide gel: Interpretation of results
The analysis of the products from the ‘plus’ and ‘minus’ reactions demands an acrylamide gel capable of resolving oligonucleotides which differ in length by only one residue. A 12% polyacrylamide slab gel in a Tris-borate buffer containing 8M-urea is usually employed for this purpose. Since the adequate resolution of products is the main limiting factor in this type of analysis and since this in turn depends on obtaining a sharp autoradiograph it is important to use extremely thin gels to avoid the blurring which would unavoidably result from the high energy p-emission of [PI embedded in a thicker gel. We define the smallest oligonucleotide in the -T channel as band 1. This means that the next residue after the 3’-terminus of the oligonucleotide corresponding to band 1 will be a T. This is equivalent to, and is confirmed by the presence of a band in the +T channel corresponding to the next longest oligonucleotide. Band 2 occurs in the +T channel and -A channel showing that its 3’-terminus is T and the adjacent nucleotide is an A, thus defining the dinucleotide sequence T-A. The next longest oligonucleotide occurs in the +A and -C channels. This defines the dinucleotide A-C and so extends the sequence to T-A-C.
Typically a sequence of 60-100 nucleotides, starting about 10-20 nucleotides from the 3’-end of the primer sequence, can be obtained from a single gel. In the above example each nucleotide in the sequence is represented by bands in both the + and – channels. However, if a run of two or more identical nucleotides occur, only the first one will be seen in the minus reaction and only the last one of the run will be seen in the equivalent plus reaction.
If a synthetic oligonucleotide or a small restriction fragment (<l00 nucleotides) is used as a primer for the initial extension, the products of the plus and minus reactions can be analysed directly. Clearly, the smaller the primer that can be used, consistent with its ability to yield a unique primer-template complex in the annealing reaction, the greater the amount of sequence information that can be deduced since the extension reactions can be pushed further and still yield resolvable fragments. When using primers of this sort it is important to maintain the integrity of the 5’-terminus so that the difference in length of the fragments depends only on differences at their 3’ termini. This is achieved by using DNA polymerase lacking the normal 5’- exonuclease activity of DNA polymerase I. If longer restriction fragments are used as primers it becomes necessary to cleave the primer from the can be deduced for the synthesized oligonucleotide. Typically a sequence of 60-100 nucleotides, starting about 10-20 nucleotides from the 3’-end of the primer sequence, can be obtained from a single gel. In the above example each nucleotide in the sequence is represented by bands in both the + and – channels. However, if a run of two or more identical nucleotides occur, only the first one will be seen in the minus reaction and only the last one of the run will be seen in the equivalent plus reaction.
The distance separating the different bands, in either the plus or minus patterns, will define how many residues there are in a run. This can lead to problems in the precise determination of the lengths of longer ’runs’ and, partly for that reason, it is advantageous to include a ‘zero’ channel on the sequencing gel. This is simply an aliquot of the initial reaction which ideally will contain labelled oligonucleotides of all possible chain lengths and will therefore yield distinct bands corresponding to each residue of a run. This is conveniently done by digestion with the restriction enzyme originally used to prepare the primer. The products for analysis in this case represent only the denovo synthesized sequences and in consequence are theoretically capable of yielding relatively more sequence data than in experiments where the primer remains attached to the analysed products. Some restriction enzymes, however, are inhibited by the single-stranded DNA present in uncopied regions of the template and these enzymes cannot therefore be reliably used to cleave the primer from the extended product. One way round this problem is to use the single-site ribosubstitution method, in which a single ribonucleotide is incorporated atthe priming site thus allowing the primer to be cleaved from the extended product with ribonuclease or alkali. This method is also useful if the restriction endonuclease used to generate the primer has a second cleavage site within the region to be sequenced.
Additional methods useful in conjunction with plus and minus method
In the protocol described above restriction fragments are used as primers for the synthesis of cDNA, and the same endonuclease is subsequently used to remove the primer and generate a unique 5′-terminus on the cDNA. However, as pointed out earlier, some restriction enzymes are strongly inhibited by the single stranded regions present in the template and cannot be used to cleave at the restriction site. An additional problem arises if a second cleavage site for the same enzyme is present within the sequence copied into the radioactive DNA. Two sets of fragments would be generated and a unique sequence would not be obtained. In the ribosubstitution method these problems are circumvented by the addition of one or more ribonucleotides between the DNA primer and the radioactive cDNA. This site is susceptible to cleavage with ribonuclease or alkali.
In the presence of Mn++ ions, E. coli DNA polymerase I will incorporate ribonucleotides into DNA. In the single site ribosubstitution reaction a ribonucleotide is incorporated at the 3’-end of a DNA primer in the presence of Mn2+, and with no other triphosphates present. Further ribonucleotide incorporation is effectively suppressed in the subsequent elongation reaction by the addition of deoxyribonucleoside triphosphates.
Step 1 : Annealing reaction
Mix: 5 µl of primer (approx 1-2 pmol in H2O), 1 µl DNA template (single strand, approx. 0.4 pmol in H2O), 1.25 ml M NaCl, 1.25 ml 10 X Pol mix and 6.5 µl H2O – Seal in glass capillary tube (approx. 10 cm long x 1 mm internal diameter). Denature by heating to 100°C for 3 min, anneal by incubating at 67°C for 45 min.
Step 2: Primed synthesis of r3’P]-cDNA
- Dry down 20 Ci, if activity about 300Ci/mMol in a siliconized tube* in vacuo. (This is conveniently done as soon as the annealing reaction is started.) Annealed reaction mixtures from step 1.
To dry dATP add:
- 5 µl dCTP (0.5 mM)
- 5 µl dlITP (0.5 mM)
- 5 µl dGTP (0.5 mM)
- 5 µl 10 x Pol mix
- Mix contents by sucking up and down in capillary from siliconised tube at 0°C.
- Start reaction by mixing in 1 µl DNA polymerase I.
- Hold at 0°C.
- Remove aliquot (approx 15 µl) after 1 min and eject into 25 µl 0.1 M EDTA, pH 7.6 to stop the reaction. After 3 min eject remainder of reaction mixture into the same EDTA.
Step 3: Removal of polymerase and triphosphates
- To the extension mixture from Step 2 add 25 l phenol
- Vortex for 0.5-1 min.
- Extract 5 times with 1 ml-portions of ether to remove phenol.
- Remove last traces of ether with a stream of air or nitrogen.
- Load the sample onto a column of G-100 Sephadex ( 3 m X 200 mm) equilibrated with a degassed buffer containing 10-4 M EDTA,5 x 10-3 M Tris-HCl, pH 7.5.
- The polynucleotide is eluted with the break-through volume of the column (flow rate approx 50pVmin). Fractions of 2-3 drops may be collected and the position of the eluted DNA located by a hand held mini-monitor. Appropriate fractions are combined and freeze-dried.
- At this stage the product should register >300c.p.s. on the mini-monitor, equivalent to an incorporation of about 5%- 10% of the radioactive nucleotide. Under these conditions the extension of the primer ranges from zero to 150 to 200 nucleotides.
Step 4 : Plus and minus reactions
- Dissolve the [32P]-labelled extended polynucleotide in 20 l H2O
- Set up 8 capillaries with drawn out tips, resting tip down in siliconized tubes, on ice. Introduce into the tip of each capillary 2 pl of the polynucleotide solution and 2p1 of the appropriate plus or minus mix.
- Add 1 p1 T4 polymerase to capillaries 1-4, mix as before and incubate at 37°C for 45 min.
- Add 1 pl ‘Klenow’ polymerase to capillaries 5-8, mix and incubate at 0°C for 45 min.
- The next step depends on whether a short (< 100 nucleotides) or longer polynucleotide was used as a primer. In the former case the reaction is terminated by blowing each reaction mixture into lop1 formamide-dye mix and the remaining radioactive polynucleotide from step 3 (approx. 4 pl) is added to lop1 formamide-dye mix. This is the ‘zero’ sample. The nine samples are now ready to proceed to step 5. Where a longer primer was used, this needs to be cleaved from the product using the appropriate restriction endonuclease. Add 1 µl, (0.5 to 1.0 unit) of the datum restriction endonuclease to each sample, mix, and incubate at 37°C for 30min. (The datum endonuclease is that which defines the 5’ of the sequence under investigation, which will usually be the endonuclease used to prepare the primer fragment.) Stop the reaction by blowing into 10 µl formamide-dye mix.
- In addition, to provide the reference pattern of oligonucleotides (the zero channel) a sample of the radioactive polynucleotide prepared in Step 3 is also digested with the restriction endonuclease.
- Mix: 2 µl radioactive polynucleotide from step 3, 1.5 µl H2O, 0.5 µl x 10 restriction buffer,1 µl restriction endonuclease (0.5 to 1.0 unit).Incubate 37″C, 30 min Stop reaction by blowing into lop1 forrnamide-dye mixture.
Step 5 : Gel electrophoresis
(i) Preparation of the acrylamide gel
This is conveniently done while the extension product is drying (step 3). The gel is cast in a cell made from two tempered glass plates,* 40 cm x 20 crn separated by two ‘perspex’ (polymethylmethacrylate sheet) spacers, 1.0-1.5 mm thick, running the length of the gel compartment. The design is essentially that of Studier (1973). The cell is sealed along the bottom and both sides with waterproof tape and immediately after pouring the gel a close-fitting well former giving 12 wells (1.1cm wide) is inserted and the gel allowed to set in the near horizontal position. Immediately before the gel is required the tape is peeled off the bottom of the cell and the well former carefully removed (care is required not to break the wells). The cell is clamped vertically in the electrophoresis apparatus and the buffer compartments filled with 1 xTBE buffer..
- Dissolve 63 g urea (AnalaR) in 15 ml 10 X TBE plus 60 ml 30%
- Add 5 ml 1.6% freshly prepared ammonium persulphate and
- Degas on water pump.
- Add 75 µl TEMED (N,N,N’,N’,-tetramethylethylenediamine). Mix gently.
- Pour gel immediately.
- The gel usually sets within 20-30min and can be used after 1 hour. Alternatively the gel can be left overnight, with the well former in place, before use.
(ii) Running the gel
Heat the nine samples from step 4 at 90” for 3min. Using a pasteur pipette blow fresh 1 x TBE into the sample wells in order to remove urea which has diffused out of the gel.
Load the samples into the gel wells using a drawn-out capillary tube.
A suitable order is:
- Run the gel at about 6OOV until the fast migrating dye (bromophenol blue) is at the bottom of the gel (approx. 4 hr). The gel gets quite hot during electrophoresis ensuring the DNA remains fully denatured.
- Remove one glass plate from the gel and cover the exposed surface with cellophane film. Label with radioactive ink (preferably 35S) and autoradiograph for 1-2 days at -20°C .Alternatively the gel may be fixed by immersion in 10% acetic acid for 15-20min, washed 1-2 min in distilled water, blotted dry with absorbent paper, covered with cellophane and autoradiographed at room temperature.
(iii) Enzymes and chemicals
Formamide-dye mix: 0.03% xylene cyanol FF, 0.03% bromophenol blue, 25 mM-EDTA in 90% formamide.
30% acrylamide: 29% (w/v) acrylamide, 1% (w/v) bis-acrylamide. Deionized by stirring with Amberlite MB-1 (5 g/100 mi) for 1 hr and filtering.
Stop mix: 0.03% bromophenol blue, 40% sucrose, 25mM EDTA in H2O.