Chapter - 12: DNA Recombinant Technology

Chapter 12 Outline

Restriction endonucleases

History

Naming the restriction enzyme

Recognition sites

Restriction-modification system(R-M system)

EcoR1 and EcoR1 methylase

Restriction maps

Cut and paste technique

Gel electrophoresis

Polyacrylamide gel electrophoresis

Agarose gel electrophoresis

Blotting techniques

Southern blotting

DNA fingerprinting

Northern blotting

S1 nuclease protection assay

DNA sequencing

Sanger’s Method

Maxam-Gilbert sequencing

Automated sequencing

Restriction Endonucleases

Restriction endonucleases are also known as restriction enzymes that protect the host by preventing the invasion of foreign DNA by cutting it up. Since these enzymes cut at sites within the foreign DNA, they are called the endonucleases. Cohen and Boyer were the first to name them restriction endonucleases and it was first discovered in E. coli in 1960 by Stewart Lynn and Werner Arber. A restriction cuts double-stranded or single stranded DNA at specific recognition nucleotide sequences known as restriction sites. The restriction enzymes are sequence specific and cut whenever these sequences occur and only when these sequences occur.

History

After isolating the first restriction enzyme, HindII, in 1970, and the subsequent discovery and characterization of numerous restriction endonucleases, the 1978 Nobel Prize for Physiology or Medicine was awarded to Daniel Nathans, Werner Arber, and Hamilton O. Smith Their discovery led to the development of recombinant DNA technology that allowed, for example, the large scale production of human insulin for diabetics using E.coli bacteria. Over 3000 restriction enzymes have been studied in detail, and more than 600 of these are available commercially and are routinely used for DNA modification and manipulation in laboratories.

Naming the Restriction Enzyme

The restriction enzymes derive the first three letters of their names from the Latin name of the microorganism that produces them. The first letter is the first letter of the genus and the next two letters are the first two letters of the species. Furthermore, the strain designation is sometimes included. If the strain produces one restriction enzyme the name ends with the roman numerical I and if more of them re produced they are numbered as II, III and so on.

Derivation of the EcoRI nameAbbreviation Meaning Description

E Escherichia genusco coli speciesR RY13 strain

I First identified order of identificationin the bacterium

Recognition Site

Recognition sites are the sequences where the restriction enzyme makes the cut. It is sometimes refered as the cutting sites. The sequence is often four to six nucleotide long such as the restriction endonucleases EcoRI recognizes the sequence GAATTC. Whenever the sequence occurs and only when this sequence occurs, the specific restriction enzymes will make the cut. Each restriction enzyme has it’s own cutting sequence. Restriction enzymes can make staggered cuts because the sequences they recognize usually display two-fold symmetry. Meaning they are identical after rotating them 180 degrees.

5'-GTATAC-3'::::::

3'-CATATG-5'

The sequences read the same forward and backward. The sequences with twofold symmetry are also called palindromes. A palindrome recognition site reads the same on the reverse strand as it does on the forward strand. Restriction enzymes always read from the five prime end to the three prime end. The two stands of the DNA duplex have the same nucleotide sequence running in opposite directions for the length of the recognition sequence. Because the same recognition sequence occurs in both stands of the DNA duplex, the restriction enzyme can bind to and cleave both strands of the DNA molecule. Because the bond cleaved is typically not positioned in the center of the recognition sequence, and the DNA strands are antiparallel, the cut sites are offset from each other. In the table below, it shows some of the common retriction enzymes and their cutting sites.

Figure 1: Restriction enzymes cut DNA molecules at specific sequences

(only 5’-->3’ is shown); arrows are cutting sites

Some of the restriction enzymes cut at 4-bp sequences rather than the 6-bp sequences. These are more common and cut more frequently. Because in a given sequence the 4-bp will occur about once every four to the power four, every 256 bp and while the 6-bp sequence will occur every four ot the power six, every 4096 bp. Some restriction enzymes are called the rare cutters such as NotI because it cuts every 8-bp sequence. Some of the restriction enzymes are called the Isoschizomers because they recognize the identical sequences but cutting sites within the sequences are different.

The main advantage off the restriction enzyme is that it can cutDNA stands reproducibly at the same places which is the basis of the techniques used to analyze genes and their expression. Many restriction enzymes make staggered cuts in the two DNA stands, leaving single-stranded overhangs called sticky ends that can base-pair together briefly. This makes it easier to switch two different DNA molecules together. EcoRI produces "sticky" ends,

Here, we get the 4-base overhangs that protrudes from the 5 prime ends of the fragments.

---G3 + 5’AATTC------CTTAA5’+3’G---

When the sequences they recognize are the two fold symmetry the restriction enzymes makes the staggered cuts.These end products are called the blunt ends. These sequences are identical after rotating them 180 degrees. SmaI restriction enzyme cleavage produces "blunt" ends

In the following table we see some of the restriction enzymes and the sticky or blunt ends produced by these enzymes. In the table below we see that the enzymes BamHI and EcoRI produce the sticky ends and the enzymes HaeIII produces the blunt ends.

Figure 2: Sticky vs. blunt ends

Restriction-modification System (R-M System)

The host cells own DNA is protected from its restriction enzyme with the Restriction-modification system. All restriction endonucleases are paired with methylases that recognize and methylate the same recognition sites. These two enzymes-the restriction endonucleases and methylases are collectivly called the Restriction-modification system. After, methylation the DNA sites rae protected against most retsriction endonucleases so the methylated DNA can persist unharmed in the host cell.

Figure 3: DNA replication during R-M modification system

This figure explains us how the DNA replicates while it is methylated. During replication, one stand of the daughter duplex will be a newly made stand and will be unmethylated. The other strand will be the parental stand and will be methylated. The half-methylation protects the DNA duplex against cleavage by the restriction enzyme. It allows the methylase time to find the newly made stand and methylate it.

EcoR1 and EcoR1 Methylase

EcoR1 does a staggered cut at a specific 6-bp inverted repeat and gives sticky ends; the corresponding methylase also recognizes the same sequence and modifies it. Cohen and Boyer took advantage of the sticky ends produced by this enzyme in their cloning experiment.

Figure 4: EcoR1 and EcoR1 methylase system

Restriction Maps

Restriction mapping is the process of obtaining structural information on a piece of DNA by the use of restriction enzymes. A restriction map is the map of known restriction sites within a sequence of DNA obtained by using the restriction enzymes. It is used as a guide to engineer the plasmids. Restriction mapping involves digesting DNA with a series of restriction enzymes and then separating the resultant DNA fragments by agarose gel electrophoresis. The distance between restriction enzyme sites can be determined by the patterns of fragments that are produced by the restriction enzyme digestion. In this way, information about the structure of an unknown piece of DNA can be obtained. One application of the restriction mapping is to cut a large piece of DNA into smaller fragments to allow it to be sequenced. Also, restriction mapping is an easy way to compare DNA fragments without having any information of their nucleotide sequence. DNA must be chopped up into smaller pieces and subcloned to perform the sequencing. For example, one can isolate two clones for a gene that are 8 kb and 10 kb long and these clones overlap since they have sequences that are common between them. A restriction map can tell you how much they overlap by:

Figure 5: Restriction mapping

From the restriction map information one can find out which parts of the two clones are identical, the parts of clones that overlap and which parts are different. In order to find out the sequence of this gene, one will only have to sequence the area of overlap in one of the clones.

Cut and Paste Technique

Restriction enzymes that cleave the DNA backbones in positions that are not directly opposite each other create overhangs. For example, EcoRI has the following recognition sequence:

Enzyme Source Recognition Sequence Cut

EcoRI Escherichia coli 5'GAATTC3'CTTAAG

5'---G AATTC---3'3'---CTTAA G---5'

When this restriction enzyme encounters this sequence, it cleaves each backbone between the G and the closest A base residues. Once the cuts have been made, the resulting fragments are held together only by the relatively weak hydrogen bonds that hold the complementary bases to each other. The weakness of these bonds allows the DNA fragments to separate from each other. Each resulting fragment has a protruding 5' end composed of unpaired bases. Other enzymes create cuts in the DNA backbone which result in protruding 3' ends. Protruding ends—both 3' and 5'—are also called "sticky ends" because they tend to bond with complementary sequences of bases. Therefore, if an unpaired length of bases (5' A A T T 3') encounters another unpaired length with the sequence (3' T T A A 5') they will bond to each other. Ligase enzyme is then used to join the phosphate backbones of the two molecules. Any pair of complementary sequences will tend to bond, even if one of the sequences comes from a length of human DNA, and the other comes from a length of bacterial DNA. This technique has allowed the production of recombinant DNA molecules.

Figure 6: The cut and paste technique of the restriction enzyme EcoRI.

Figure 7: The Ligase reaction

The mechanism of DNA ligase is to form two covalent phosphodiester bonds between 3' hydroxyl ends of one nucleotide with the 5' phosphate end of another. ATP is required for the ligase reaction. The reaction occurs in three stages in all DNA ligases:

1.      Formation of a covalent enzyme-AMP intermediate linked to a lysine side-chain in the enzyme.

2.      Transfer of the AMP nucleotide to the 5’ phosphate of the nicked DNA strand.

3. Attack on the AMP-DNA bond by the 3’-OH of the nicked DNA sealing the phosphate backbone and resealing AMP.

Gel ElectrophoresisGel electrophoresis is a technique used in the laboratory that results in the separation of DNA, RNA or protein molecules using an electric field applied to a gel matrix. It s usually applied to DNA after PCR. However, for characterization of DNA, it is often used before any of the methods.

Figure 8: Gel electrophoresis apparatus

The gel used for this method is crosslinked polymer whose composition and porosity is chosen based on the specific weight and composition of the target molecules that the gel contains and separates. To separate the small molecules the gel contains different concentrations of acrylamide and a cross-linker and when separating large molecules the gel contains purified agarose. "Electrophoresis" refers to the electromotive force (EMF) that is used to move the molecules through the gel matrix. By placing the molecules in wells in the gel and applying an electric field, the molecules will move through the matrix at different rates, determined largely by their mass when the charge to mass ratio (Z) of all species is uniform, toward the anode if negatively charged or toward the cathode if positively charged.

After the electrophoresis is complete, the molecules in the gel can be stained with dyes like ethidium bromide, silver, or coomassie blue dye to make them visible. If the analyte molecules fluoresce under ultraviolet light, a photograph can be taken of the gel under ultraviolet lighting conditions. If the molecules to be separated contain radioactivity added for visibility, an autoradiogram can be recorded of the gel. If several mixtures have initially been injected next to each other, they will run parallel in individual lanes. Depending on the number of different molecules, each lane shows separation of the components from the original mixture as one or more distinct bands, one band per component. Incomplete separation of the components can lead to overlapping bands, or to indistinguishable smears representing multiple unresolved components. Bands in different lanes that end up at the same distance from the top contain molecules that passed through the gel with the same speed, which usually means they are approximately the same size. If such a marker was run on one lane in the gel parallel to the unknown samples, the bands observed can be compared to those of the unknown in order to determine their size.

Figure 9: An example of the gel electrophoreses result.

Gel electrophoresis is used in forensics, molecular biology, genetics, microbiology and biochemistry. The image is recorded with a computer operated camera, and the intensity of the band or spot of interest is measured and compared against standard or markers loaded on the same gel. The measurement and analysis are mostly done with specialized software. In the case of nucleic acids, the direction of migration, from negative to positive electrodes, is due to the naturally-occurring negative charge carried by their sugar-phosphate backbone.

Polyacrylamide Gel ElectrophoresisThis technique is widely used in to separate proteins according to their electrophoretic mobility (a function of length of polypeptide chain or molecular weight). SDS-PAGE is an abbreviation for sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis.

Figure 10: Polyacrylamide gel electrophoresis

In what is probably the most powerful technique for resolving protein mixtures, proteins are exposed to ionic detergent SDS (sodium dodecylsulfate) before and during gel electrophoresis.  SDS denatures proteins, causing multimric proteins to dissociate into their subunits, and all polypeptide chains are forced into extended conformations with similar charge:mass ratios.  SDS treatment therefore eliminates the effects of differences in shape so that chain length, which

reflects mass, is the sole determinant of the migration rate of proteins in SDS- polyacrylamide electrophoresis.

Agarose Gel Electrophoresis

This method is used to separate DNA, RNA or protein. This method uses the sizes of the molecules to separate them. This is achieved by moving negatively charged nucleic acid molecules through an agarose matrix with an electric field shorter molecules move faster and migrate farther than longer ones because the sieving effect of the gel. Proteins are separated by charge in this method. This method is used to estimate the size of DNA molecules following restriction enzyme digestion, analysis of the products from PCR, separation of restricted genomic DNA. The advantages are that the gel is easily poured, does not denature the samples. The samples can also be recovered. The disadvantages are that gels can melt during electrophoresis, the buffer can become exhausted, and different forms of genetic material may run in unpredictable forms. After the experiment is finished, the resulting gel can be stored in a plastic bag in a refrigerator.

The most important factor is the length of the DNA molecule, smaller molecules travel farther. Increasing the agarose concentration of a gel reduces the migration speed and enables separation of smaller DNA molecules. The higher the voltage, the faster the DNA moves. Conformations of a DNA plasmid that has not been cut with a restriction enzyme will move with different speeds (slowest to fastest: nicked or open circular, linearised, or supercoiled plasmid).

Figure 11: Agarose gel electrophoresis

The image to the right shows migration of a set of DNA fragments in three concentrations of agarose, all of which were in the same gel tray and electrophoresed at the same voltage and for identical times. Notice how the larger fragments are much better resolved in the 0.7% gel, while

the small fragments separated best in 1.5% agarose. The 1000 bp fragment is indicated in each lane

The most common dye used to make DNA or RNA bands visible for agarose gel electrophoresis is ethidium bromide which fluoresces under UV light when intercalated into DNA (or RNA). Since the stained DNA is not visible in natural light, scientists mix DNA with negatively charged loading buffers such as Bromophenol blue before adding the mixture to the gel.

Blotting techniqueBlotting techniques are used to transfer DNA, RNA and proteins into a carrier so that they can be separated flowing gel elctrophorises. The southern blot is used for DNA, the northern blot is used for RNA and western blot for proteins. A specific DNA sequence isolated by cloning can serve as a probe to detect the presence and the amounts of complementary nucleic acids in complex mixtures including total cellular DNA or RNA.

Figure 12: Blotting technique

The figure above illustrates the steps in the blotting technique-dsDNA denature or melt bind onto filter paper, incubate with labeld ss DNA probe, complementary DNA hybridizes, wash away non-bound labeled DNA, perform autoradiography.

Figure 13: Detecting nucleic acid with a non-radioactive probe

The above figure shows the steps in detecting nucleic acid with a non-radioactive probe-replicate the probe DNA in the presence of dUTP-biotin, denature, hybridize with target DNA, mix with bifunctional reagent avidin-AP, alkaline phosphatase can cleave the P-reagent and generate chemiluminescence for autoradiographic detection.

Southern BlottingThe Southern blotting is a method routinely used to detect specific DNA sequence in a complex DNA samples. The method is named after its inventor Edwin Southern in 1970. It combines electrophoresis-separated DNA fragments to a filter membrane and subsequent fragment detection by probe hybridization. It has been applied to detect Restriction Fragment Length Polymorphism (RFLP) and Variable Number of Tandem Repeat Polymorphism (VNTR).  The latter is the basis of DNA fingerprinting.

Figure 14: Southern blotting

The figure above shows the steps in Southern blotting.  (a) The DNA to be analyzed is digested with restriction enzymes and then separated by agarose gel electrophoresis.  (b) The DNA fragments in the gel are denatured with alkaline solution and transferred onto a nitrocellulose filter or nylon membrane by blotting, preserving the distribution of the DNA fragments in the gel.  (c) The nitrocellulose filter is incubated with a specific probe.  The location of the DNA fragment that hybridizes with the probe can be displayed by autoradiography. Autoradiography is a technique used in southern blotting where a radioactive sample is exposed to a photographic emulsion, thus “taking a picture of itself”.

DNA FingerprintingDNA fingerprinting is a used to assist in the identification of individuals on the basis of their respective DNA profiles. DNA profiles are encrypted sets of numbers that reflect a person’s DNA makeup, which can also be used as the person’s identifier. Even though the chemical structure of everyone’s DNA is the same, there are differences between the orders of the base pairs. There are so many millions of base pairs in each person’s DNA that every person has a different sequence. Using these sequences, every person could be identified solely by the sequence of their base pairs. Because there are so many millions of base pairs, the task would be very time-consuming. However, scientists are able to use a shorter method, because of repeating patterns in DNA. DNA profiling uses repetitive (“repeat”) sequences that are highly variable, called variable number tandem repeats (VNTR). VNTRs loci are very similar between closely related humans, but so variable that unrelated individuals are extremely unlikely to have the same VNTRs. Sir Alec Jeffreys at the University of Leicester in England reported it first in 1984. It is used in, for example, parental testing and rape investigation.

Figure 15: DNA fingerprinting

In this figure, cut DNA with HaeIII into 8 fragments, only 3 contains minisatellites; electrophoresis, denature, Southern blot, hybridize with radioactive probe.

DNA fingerprinting is essentially an application of southern blotting. The frequency of a coincidental match of the DNA pattern between two individuals is one in 33 billion.

Northern BlottingA Northern blot is a blot techniques used to transfer RNA (or isolated mRNA) onto a carrier for sorting and identification to study gene expression. Northern blots can be used to determine the size of an mRNA transcript, identify functional variants of a gene and measure the relatedness of species. It also provides a direct relative comparison of message abundance between samples on a single membrane. It is the preferred method for determining transcript size and for detecting alternatively spliced transcripts. The Northern blot takes its name from its predecessor, the Southern blot, which was named for biologist Edwin Southern. The northern blot technique was developed in 1977 by James Alwine, David Kemp, and George Stark at Stanford University. The Northern blotting procedure is straightforward and provides opportunities to evaluate progress at various points (e.g., integrity of the RNA sample and how efficiently it has transferred to the membrane).

RNA samples are first separated by size via electrophoresis in an agarose gel under denaturing conditions. A blotting membrane is applied to a gel electrophoreses to get the molecules to transfer from the gel to the membrane. The gel is treated with a formaldehyde hybridization buffer to denature the RNA. The membrane should also be kept moist throughout. After pressure is applied evenly to the membrane, the RNA migrates and sticks to the membrane. Then the membrane is baked or irradiated, permanently attaching the RNA to the membrane. The RNA is then transferred to a membrane, crosslinked and hybridized with a labeled probe. A hybridization probe (known-sequence single-strand DNA or RNA designed to match the analyzed RNA) is introduced to pair with the blotted DNA sequences; the probe is tagged with fluorescent or chromogenic dyes for identification. Northern hybridization is exceptionally versatile in that radiolabeled or nonisotopically labeled DNA, in vitro transcribed RNA and oligonucleotides can all be used as hybridization probes.

Figure 16: Flow diagram outlining the general procedure for RNA detection by northern blotting.

The steps involved in Northern analysis include:

RNA isolation (total or poly(A) RNA) Probe generation

Denaturing agarose gel electrophoresis

Transfer to solid support and immobilization

Prehybridization and hybridization with probe

Washing

Detection

Northern blotting allows one to observe a particular gene's expression pattern between tissues, organs, developmental stages, environmental stress levels, pathogen infection, and over the course of treatment. The technique has been used to show overexpression of oncogenes and downregulation of tumor-suppressor genes in cancerous cells when compared to 'normal' tissue, as well as the gene expression in the rejection of transplanted organs. Despite these advantages, there are limitations associated with Northern analysis such as samples are even slightly degraded, is, in general, less sensitive than nuclease protection assays, and the difficulty associated with multiple probe analysis.

S1 Nuclease Protection AssayNuclease protection assays are an extremely sensitive method for the detection, quantitation and mapping of specific RNAs in a complex mixture of total cellular RNA. The technique can identify one or more RNA molecules of known sequence even at low total concentration. The extracted RNA is first mixed with antisense RNA or DNA probes that are complementary to the sequence or sequences of interest and the complementary strands are hybridize to form double-stranded RNA (or a DNA-RNA hybrid).After hybridization, any remaining unhybridized probe and sample RNA are removed by digestion with a mixture of nucleases that specifically cleave only single-stranded RNA but have no activity against double-stranded RNA. These products are separated on a denaturing polyacrylamide gel and are visualized by autoradiography.

Figure 17: Detection of Specific mRNA Species Using a Nuclease Protection Assay.

When the reaction runs to completion, susceptible RNA regions are degraded to very short oligomres or to individual nucleotides; the surviving RNA fragments are those that were complementary to the added antisense strand and thus contained the sequence of interest. When the probe is a DNA molecule, S1 nuclease is used; when the probe is RNA, any single-strand-specific ribonuclease can be used. Thus the surviving probe-mRNA complement is simply detected by autoradiography.

Figure 18: Nuclease protection method for quantitating specific RNAs in a mixture and mapping them.

Nuclease protection assays are used to map introns and 5' and 3'; ends of transcribed gene regions. Quantitative results can be obtained regarding the amount of the target RNA present in the original cellular extract - if the target is a messenger RNA, this can indicate the level of transcription of the gene in the cell. Nuclease protection assays are more sensitive, more tolerant of partially degraded RNA, are able to distinguish between transcripts of multi-gene families, can be used to map mRNA and Multi-probe assays are easy to perform with. However, the primary limitation of NPAs is the lack of information on transcript size, and the lack of probe flexibility.

DNA Sequencing

DNA sequencing refers to sequencing methods which determine the order of the nucleotide bases—adenine(A), guanine(G), cytosine(C), and thymine(T)—in a molecule of DNA.

DNA sequencing has become the most important tool in numerous applied fields such as diagnostic, biotechnology, forensic biology and biological systematics. The rapid spped in attaining the modern DNA sequening technology has been instrumental in the sequencing of the human genome, in the Human Genome Project.

Figure 19: DNA sequencing

The first DNA sequences were obtained in the early 1970. RNA sequencing was one of the earliest forms of nucleotide sequencing. Walter Fiers and his coworkers at the University of Ghent (Ghent, Belgium), was the first to complete the genome of Bacteriophage MS2 between 1972 and 1976. In the early 1970s by Frederick Sanger at the University of Cambridge, in England and Walter Gilbert and Allan Maxam at Harvard, used a number of laborious methods to sequencing. For instance, in 1973, Gilbert and Maxam reported the sequence of 24 base pairs using a method known as wandering-spot analysis. DNA sequencing has been applied to

many areas of research such as the polymerase chain reaction (PCR), a method which rapidly produces numerous copies of a desired piece of DNA and to identify restriction sites in plasmids

Sanger’s MethodDideoxynucleotide sequencing represents only one method of sequencing DNA, which is also called the Sanger’s method since Sanger devised the method. In 1945, Frederick Sanger described its use for determining the N-terminal amino acid in polypeptide chains, in particular insulin. This technique utilizes 2',3'-dideoxynucleotide triphospates (ddNTPs), molecules that differ from deoxynucleotides by the having a hydrogen atom attached to the 3' carbon rather than an OH group. These molecules terminate DNA chain elongation because they cannot form a phosphodiester bond with the next deoxynucleotide. Dinitrofluorobenzene reacts with the amine group in amino acids to produce dinitrophenyl-amino acids. These DNP-amino acids are moderately stable under acid hydrolysis conditions that break peptide bonds. The DNP-amino acids can then be recovered, and the identity of those amino acids can be discovered through chromatography.

Figure 20

Since it is more efficient and uses fewer toxic chemicals and lower amounts of radioactivity than other methods, it rapidly became the method of choice. The classical chain-termination method requires a single-stranded DNA template, a DNA primer, DNA polymerase, radioactively or fluorescently labeled nucleotides, and modified nucleotides that terminate DNA strand elongation.

Figure 21: Sanger’s method

The DNA sample is divided into four separate sequencing reactions, containing all four of the standard deoxynucleotides (dATP, dGTP, dCTP and dTTP) and the DNA polymerase. To each reaction is added only one of the four deoxynucleotides (ddATP, ddGTP, ddCTP, or ddTTP) which are the chain-terminating nucleotides, lacking an 3'-OH group required for the formation of a phosphodiester bond between two nucleotides, thus terminating DNA strand extension and resulting in DNA fragments of varying length. The newly synthesized and labeled DNA fragments are heat denatured and separated by size by gel electrophoresis on a denaturing polyacrymide-urea gel with each of the four reactions run in one of four individual lanes (lanes A, T, G, C); the DNA bands are then visualized by autoradiography or UV light, and the DNA sequence can be directly read off the X-ray film or gel image.

CAAAAAACGG... 21 Primer-5’-ATGATACGGTCT-3’

Template: 5’-AGACCGTATCAT-3

The bottom-most band indicates that its particular dideoxynucleotide was added first to the labeled primer. In the figure below, the band that migrated the farthest was in the ddATP reaction mixture. Therefore, ddATP must have been added first to the primer, and its complementary base, thymine, must have been the base present on the 3' end of the sequenced strand. One can continue reading in this fashion. If one reads the bases from the bottom up, one is reading the 5' to 3' sequence of the strand complementary to the sequenced strand. The sequenced strand can be read 5' to 3' by reading top to bottom the bases complementary to those on the gel.

Figure 21: Results of the gel electrophoresis

This figure is a representation of an acrylamide sequencing gel. The sequence of the strand of DNA complementary to the sequenced strand is 5' to 3' ACGCCCGAGTAGCCCAGATT while the sequence of the sequenced strand, 5' to 3', is AATCTGGGCTACTCGGGCGT.

Figure 22: An example of Sanger’s method in action. Sanger sequencing: the reading 5’-ATGATACGGTCT from the primer extension

Maxam-Gilbert SequencingThe Maxam-Gilbert method of nucleotide sequence determination is based on preferential, base-specific methylation followed by chemical cleavage to generate a nested set of end- labeled derivatives. In 1976-1977, Allan Maxam and Walter Gilbert, two years after the

discovery of the Sanger’s method, developed a DNA sequencing method based on chemical modification of DNA and subsequent cleavage at specific bases. Maxam-Gilbert sequencing rapidly became more popular, since purified DNA could be used directly, while the initial Sanger method required that each read start be cloned for production of single-stranded DNA. However, it has fallen out of favor due to its technical complexity prohibiting its use in standard molecular biology kits, extensive use of hazardous chemicals, and difficulties with scale-up. With this method of DNA sequencing instead of synthesizing DNA in vitro and stopping the synthesis reactions with chain terminators, this method starts with full-length, end labeled DNA and cleaves it with base specific reagents.

The method requires radioactive labeling at one end and purification of the DNA fragment to be sequenced. Chemical treatment generates breaks at a small proportion of one or two of the four nucleotide bases in each of four reactions (G, A+G, C, C+T). Thus a series of labeled fragments is generated, from the radiolabelled end to the first 'cut' site in each molecule. The fragments in the four reactions are arranged side by side in gel electrophoresis for size separation. To visualize the fragments, the gel is exposed to X-ray film for autoradiography, yielding a series of dark bands each corresponding to a radiolabelled DNA fragment, from which the sequence may be inferred.

Figure 23: DNA sequencing with Maxam-Gilbert method

Maxam-Gilbert Method: Breaking the end-labeled DNA strand at specific bases using base-specific reagents. (i) radiolabel, methylation with dimethyl sulfate, treat with piperidine, remove base, break DNA at the apurinic site, leaves 3’-phosphate on the nucleotide preceded the G. All G-removed fragments can be separated by gel.

It is also known as 'chemical sequencing', this method originated in the study of DNA-protein interactions (footprinting), nucleic acid structure and epigenetic modifications to DNA, and within these it still has important applications. 

Figure 24: The Steps of Maxam-Gilbert method

This can be 5’- or 3’-end labeling. Next, we modify one kind of base. Here we use dimethyl sulfate (DMS) to methylate guanines. (Actually, this reagent also methylates adenines, but not in a way that leads to DNA strand cleavage.) As in the chain termination method, we do not want to affect every guanine, or we will produce only tiny fragments that will not allow us to determine the DNA’s sequence. Therefore, we do the methylation under mild conditions that lead to an average of only one methylated guanine per DNA strand. Next, we use a reagent (piperidine) that does two things: it causes loss of the methylated base, then it breaks the DNA backbone at the site of the lost base (the apurinic site). In this case, the G in the middle of the sequence was methylated, so strand breakage occurred there, producing a labeled trimer.

Figure 25: An example of the results obtained form Maxam-Gilbert method

The reading of the sequence 5’-CTTTTTTGGGCTTAGC…., gives a direct reading of the sequence upto 500 nt.

Automated SequencingAutomated DNA-sequencing is a technique that can sequence up to 384 DNA samples in a single batch in up to 24 runs a day. DNA sequencers carry out capillary electrophoresis for size separation, detection and recording of dye fluorescence, and data output as fluorescent peak trace chromatograms. Sequencing reactions by thermocycling, cleanup and re-suspension in a buffer solution before loading onto the sequencer are performed separately. A number of commercial and non-commercial software packages can trim low-quality DNA traces automatically. These programs score the quality of each peak and remove low-quality base peaks (generally located at the ends of the sequence). The accuracy of such algorithms is below visual examination by a human operator, but sufficient for automated processing of large sequence data sets.

Figure 26: Automated Sequencing

In the figure above, we see the steps for automated sequencing and below is the graph produced by this method.

References

http://www.biochem.umd.edu/biochem/kahn/molmachines/replication/DNA%20Ligase.htm

http://faculty.plattsburgh.edu/donald.slish/Restmap.html

http://www.bio.davidson.edu/courses/Bio111/seq.html

http://www.dnasequencing.org/maxam-gilbert

www.wikipedia.org

http://www.web-books.com/MoBio/Free/Ch9D.htm

http://www.molecularstation.com/sds-page-gel-electrophoresis/#definition

http://www.molecularstation.com/images/southern-blot.jpg

http://users.rcn.com/jkimball.ma.ultranet/BiologyPages/D/DNAsequencing.html

http://www.molecularstation.com/images/northern-blot-med.jpg

http://www.ambion.com/techlib/basics/northerns/index.html

http://www.iscid.org/encyclopedia/Northern_Blot

http://www.ambion.com/techlib/basics/npa/index.html