DNA footprinting

From Wikipedia, the free encyclopedia

DNA footprinting is a method of investigating the sequence specificity of DNA-binding proteins. This technique can be used to study protein-DNA interactions both outside and within cells.

The regulation of transcription has been studied extensively, and yet there is still so much that is not known. Transcription factors and associated proteins that bind promoters, enhancers, or silencers to drive or repress transcription are fundamental to understanding the unique regulation of individual genes within the genome. Techniques like DNA footprinting will help elucidate which proteins bind to these regions of DNA and unravel the complexities of transcriptional control.Contents [hide]

1 Method

1.1 Labeling

1.2 Cleavage agent

2 Advanced Applications

2.1 In vivo footprinting

2.2 Quantitative footprinting

3 History

4 References

[edit]

Method

Figure 1. DNA footprinting workflow

The simplest application of this technique is to assess whether a given protein binds to a region of interest within a DNA molecule. The wet lab methodology is summarized, with appropriate selection of reagents discussed, below.[1]

Polymerase chain reaction (PCR) amplify and label region of interest that contains a potential protein-binding site, ideally amplicon is between 50 to 200 base pairs in length.

Add protein of interest to a portion of the labeled template DNA; a portion should remain separate without protein, for later comparison

Add a cleavage agent to both portions of DNA template. The cleavage agent is a chemical or enzyme that will cut at random locations in a sequence independent manner. The reaction should occur just long enough to cut each DNA molecule in only one location. A protein that specifically binds a region within the DNA template will protect the DNA it is bound to from the cleavage agent.

Run both samples side by side on a polyacrylamide gel electrophoresis. The portion of DNA template without protein will be cut at random locations, and thus when it is run on a gel, will produce a ladder-like distribution. The DNA template with the protein will result in ladder distribution with a break in it, the "footprint", where the DNA has been protected from the cleavage agent.

Note: Maxam-Gilbert chemical DNA sequencing can be run alongside the samples on the polyacrylamide gel to allow the prediction of the exact location of ligand binding site.

[edit]

Labeling

The DNA template can be labeled at the 3' or 5' end, depending on the location of the binding site(s). Labels that can be used are:

Radioactivity has been traditionally used to label DNA fragments for footprinting analysis, as the method was originally developed from the Maxam-Gilbert chemical sequencing technique. Radioactive labeling is very sensitive and is optimal for visualizing small amounts of DNA.

Fluorescence is a desirable advancement due to the hazards of using radio-chemicals. However, it has been more difficult to optimize because it is not always sensitive enough to detect the low concentrations of the target DNA strands used in DNA footprinting experiments. Electrophoretic sequencing gels or capillary electrophoresis have been successful in analyzing footprinting of fluorescently tagged fragments. [1]

[edit]

Cleavage agent

A variety of cleavage agents can be chosen. Ideally a desirable agent is one that is sequence neutral, easy to use, and is easy to control. Unfortunately none available meet all these all of these standards, so an appropriate agent can be chosen, depending on your DNA sequence and ligand of interest. The following cleavage agents are described in detail:

DNase I: a large protein that functions as a double-strand endonuclease. It binds the minor groove of DNA and cleaves the phosphodiester backbone. It is a good cleavage agent for footprinting because

its size makes it easily physically hindered. Thus is more likely to have its action blocked by a bound protein on a DNA sequence. In addition, the DNase I enzyme is easily controlled by adding EDTA to stop the reaction. There are however some limitations in using DNase I. The enzyme does not cut DNA randomly; its activity is affected by local DNA structure and sequence and therefore results in an uneven ladder. This can limit the precision of predicting a protein’s binding site on the DNA molecule. [1] [2]

Hydroxyl radicals: are created from the Fenton reaction, which involves reducing Fe2+ with H2O2 to form free hydroxyl molecules. These hydroxyl molecules react with the DNA backbone, resulting in a break. Due to their small size, the resulting DNA footprint has high resolution. Unlike DNase I they have no sequence dependence and result in a much more evenly distributed ladder. The negative aspect of using hydroxyl radicals is that they are more time consuming to use, due to a slower reaction and digestion time. [3]

Ultraviolet irradiation: can be used to excite nucleic acids and create photoreactions, which results in damaged bases in the DNA strand. Photoreactions can include: single strand breaks, interactions between or within DNA strands, reactions with solvents, or crosslinks with proteins.

The workflow for this method has an additional step, once both your protected and unprotected DNA have been treated, there is subsequent primer extension of the cleaved products. The extension will terminate upon reaching a damaged base, and thus when the PCR products are run side-by-side on a gel; the protected sample will show an additional band where the DNA was crosslinked with a bound protein.

Advantages of using UV are that it reacts very quickly and can therefore capture interactions that are only momentary. Additionally it can be applied to in vivo experiments, because UV can penetrate cell membranes. A disadvantage is that the gel can be difficult to interpret, as the bound protein does not protect the DNA, it merely alters the photoreactions in the vicinity. [4]

[edit]

Advanced Applications

[edit]

In vivo footprinting

In vivo footprinting is a technique used to analyze the protein-DNA interactions that are occurring in a cell at a given time point. DNase I can be used as a cleavage agent if the cellular membrane has been permeabilized. However the most common cleavage agent used is UV irradiation because it penetrates the cell membrane without disrupting cell state and can thus capture interactions that are sensitive to cellular changes. Once the DNA has been cleaved or damaged by UV, the cells can be lysed and DNA purified for analysis of a region of interest.

Ligation-mediated PCR is an alternative method to footprint in vivo. Once a cleavage agent has been used on the genomic DNA, resulting in single strand breaks, and the DNA is isolated, a linker is added on to the break points. A region of interest is amplified between the linker and a gene-specific primer, and when run on a polyacrylamide gel, will have a footprint where a protein was bound. [5]

In vivo footprinting combined with immunoprecipitation can be used to assess protein specificity at many locations throughout the genome. The DNA bound to a protein of interest can be immunoprecipitated with an antibody to that protein, and then specific region binding can be assessed using the DNA footprinting technique. [6]

[edit]

Quantitative footprinting

The DNA footprinting technique can be modified to assess the binding strength of a protein to a region of DNA. Using varying concentrations of the protein for the footprinting experiment, the appearance of the footprint can be observed as the concentrations increase and the proteins binding affinity can then be estimated. [1]

[edit]

History

In 1978, David Galas and Albert Schmitz developed the DNA footprinting technique to study the binding specificity of the lac repressor protein. It was originally a modification of the Maxam-Gilbert chemical sequencing technique. [7].

DNA Footprinting was developed in 1977 and is an analytical procedure in molecular biology for identifying the specific sequence of DNA (the binding site) that binds to a particular protein. DNA Footprinting is most commonly performed on proteins that are thought to play some significant functional role such as gene regulation. This method can be performed on proteins which bind both double and single-stranded DNA. Additionally, DNA-binding proteins can be split into two groups, namely site-specific DNA-binding proteins and non-specific DNA–binding proteins.

DNA Footprinting uses a damaging agent such as a chemical reagent, radical or a nuclease that can cut or modify DNA at every base pair. However, where the ligand binds to DNA, the cleavage is restrained. DNA Footprinting discovers which specific parts of a DNA molecule have sites for specific proteins to attach to them. Using this technique, DNA that has first been in the presence of DNA-binding proteins and then exposed to a damaging agent, can be compared to DNA that was never exposed to the binding protein (and thus not protected against the damaging agent). The DNA sequence that is protected from cleaving can then be identified as the binding site.

DNA Footprinting can provide information that is, conceptually, much like fingerprinting in the sense that it can be used to identify a unique individual. DNA Footprinting can extract a banding pattern, or electropherogram, much like a bar code, that can identify a species or individual (some genes will be vary at the species level and others at the individual level).

DNA Technology - What is DNA Footprinting?

Article by Rishi Prakash (2,738 pts )

Edited & published by Paul Arnold (16,776 pts ) on Aug 14, 2009

1 comment See More About: Dna Molecule

With diverse applications in the fields of biology and medical diagnostics, DNA footprinting is also a boon for researchers. It is used to detect DNA sequences to which DNA-binding proteins bind. In the following article you'll find more about the technique and its uses.

Introduction

Before tackling the topic of DNA footprinting, we need to look at some essential aspects of RNA synthesis i.e. transcription.

When DNA is converted into RNA, the process is called DNA transcription. Initiation of RNA synthesis does not take place at random points in a DNA molecule. Instead, the RNA polymerase enzyme binds to specific DNA sequences, called promoters, which direct the transcription of adjacent segments of DNA. The DNA sequences where RNA polymerases bind can be quite variable.

A process that provides information about the interface between RNA polymerase and promoters is called DNA footprinting. It is a technique that identifies the DNA sequences bound by particular proteins.

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DNA Footprinting Procedure

A DNA fragment thought to contain sequences recognized by a DNA-binding protein is isolated and radio labeled at one end of one strand.

Once the DNA fragments are labeled, they are mixed in a test tube with a DNA-binding protein and DNAse.

In a separate test tube, same labeled DNA with DNAse is mixed without the DNA-binding protein. This is for comparison.

DNAse is a cleavage agent which cuts the DNA in both samples. No cuts are made in the area where RNA polymerase has bound. It has protected the DNA.

Only labeled strands are detected in next step.

Separation of fragments is done by polyacrylamide gel electrophoresis, and visualized by autoradiogram.

Missing bands (hole or footprint) indicate where RNA polymerase was bound to DNA.

Binding of proteins to DNA sequences both near to and distant from the promoter can also affect levels of gene expression. Protein binding activates transcription by facilitating RNA polymerase binding or facilitating steps further along in the initiation process or it can repress transcription by blocking the activity of the polymerase.

DNA footprinting helps identify the promoter region and thereby the gene expression level.

DNA Footprinting Applications in Research

DNA footprinting is often used to identify the binding sites of proteins in a DNA molecule. Researchers often use this technique to identify whether a particular protein can activate or inhibit transcription. In addition, scientists also use this method to detect where proteins bind to DNA in a living cell. In this kind of experiment, scientists often grow cells under artificial conditions where the protein of interest would be more likely to bind to DNA. The resulting DNA-protein complexes are then purified from the cell, and the DNA sequences are identified.

Since DNA footprinting is being used to identify DNA sequences where proteins bind to DNA, it is of great importance in genetic research, especially in work coming from the Human Genome Project. It enables researchers to identify many functional genes present in large human DNA sequences.

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Footprinting

Footprinting is a method for determining the exact DNA sequence to which a particular DNA-binding protein binds. Examples:

hormone-receptor complexes that bind to their hormone response elements

transcription factors that bind eukaryotic operators, enhancers, and silencers

the lac repressor that shuts down the lac operon in E. coli [Discussion of the lac operon]

How, for example, does one determine the DNA sequence to which the lac repressor binds?

The procedure:

Clone a piece of DNA that contains the operator site to which the repressor binds.

Label one end of the DNA molecules with a radioactive molecule, e.g. radioactive ATP.

Digest the DNA with DNase I.

DNase I cuts DNA molecules randomly (in contrast to restriction enzymes that cut where they find a particular sequence)

Choose such gentle conditions that most molecules will be cut only once.

The result will be a mixture of radioactive fragments of varying length, with the smallest increment in length represented by a single nucleotide.

Separate the fragments by electrophoresis.

Binding of the lac repressor to the sequence of 24 base pairs in the operator prevents DNase I from attacking that region of the molecule.

When the fragments are separated by electrophoresis, those representing the lengths covered by the repressor will be missing from the autoradiogram.

The resulting gap is the "footprint".

The same sample of DNA (unprotected by the repressor) is subjected to normal DNA sequencing and the resulting ladder aligned with the footprint autoradiogram.

The exact sequence of bases in the lac operator can then be read directly because they represent the rungs of the ladder missing in the footprint.

DNA sequencing

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The term DNA sequencing refers to sequencing methods for determining the order of the nucleotide bases—adenine, guanine, cytosine, and thymine—in a molecule of DNA.

Knowledge of DNA sequences has become indispensable for basic biological research, other research branches utilizing DNA sequencing, and in numerous applied fields such as diagnostic, biotechnology, forensic biology and biological systematics. The advent of DNA sequencing has significantly accelerated biological research and discovery. The rapid speed of sequencing attained with modern DNA sequencing technology has been instrumental in the sequencing of the human genome, in the Human Genome Project. Related projects, often by scientific collaboration across continents, have generated the complete DNA sequences of many animal, plant, and microbial genomes.

The first DNA sequences were obtained in the early 1970s by academic researchers using laborious methods based on two-dimensional chromatography. Following the development of dye-based sequencing methods with automated analysis,[1] DNA sequencing has become easier and orders of magnitude faster.[2]Contents [hide]

1 History

2 Maxam–Gilbert sequencing

3 Chain-termination methods

3.1 Dye-terminator sequencing

3.2 Challenges

3.3 Automation and sample preparation

4 Amplification and clonal selection

5 High-throughput sequencing

5.1 454 pyrosequencing

5.2 Solexa sequencing

5.3 SOLiD sequencing

6 Future methods

7 Major landmarks in DNA sequencing

8 See also

9 References

[edit]

History

RNA sequencing was one of the earliest forms of nucleotide sequencing. The major landmark of RNA sequencing is the sequence of the first complete gene and the complete genome of Bacteriophage MS2, identified and published by Walter Fiers and his coworkers at the University of Ghent (Ghent, Belgium), between 1972[3] and 1976.[4]

Prior to the development of rapid DNA sequencing methods in the early 1970s by Frederick Sanger at the University of Cambridge, in England and Walter Gilbert and Allan Maxam at Harvard,[5][6] a number of laborious methods were used. For instance, in 1973, Gilbert and Maxam reported the sequence of 24 basepairs using a method known as wandering-spot analysis.[7]

The chain-termination method developed by Sanger and coworkers in 1975 soon became the method of choice, owing to its relative ease and reliability.[8][9]

[edit]

Maxam–Gilbert sequencing

In 1976–1977, Allan Maxam and Walter Gilbert developed a DNA sequencing method based on chemical modification of DNA and subsequent cleavage at specific bases.[5] Although Maxam and Gilbert published their chemical sequencing method two years after the ground-breaking paper of Sanger and Coulson on plus-minus sequencing,[8][10] 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, with the improvement of the chain-termination method (see below), Maxam-Gilbert sequencing has fallen out of favour due to its technical complexity prohibiting its use in standard molecular biology kits, extensive use of hazardous chemicals, and difficulties with scale-up.

The method requires radioactive labelling 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 labelled 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.

Also sometimes 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.

[edit]

Chain-termination methods

Part of a radioactively labelled sequencing gel

Because the chain-terminator method (or Sanger method after its developer Frederick Sanger) is more efficient and uses fewer toxic chemicals and lower amounts of radioactivity than the method of Maxam and Gilbert, it rapidly became the method of choice. The key principle of the Sanger method was the use of dideoxynucleotide triphosphates (ddNTPs) as DNA chain terminators.

The classical chain-termination method requires a single-stranded DNA template, a DNA primer, a DNA polymerase, radioactively or fluorescently labeled nucleotides, and modified nucleotides that terminate DNA strand elongation. 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 dideoxynucleotides (ddATP, ddGTP, ddCTP, or ddTTP) which are the chain-terminating nucleotides, lacking a 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 (with a resolution of just one nucleotide) by gel electrophoresis on a denaturing polyacrylamide-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. In the image on the right, X-ray film was exposed to the gel, and the dark bands correspond to DNA fragments of different lengths. A dark band in a lane indicates a DNA fragment that is the result of chain termination after incorporation of a dideoxynucleotide (ddATP, ddGTP, ddCTP, or ddTTP). The relative positions of the different bands among the four lanes are then used to read (from bottom to top) the DNA sequence.

DNA fragments are labeled with a radioactive or fluorescent tag on the primer (1), in the new DNA strand with a labeled dNTP, or with a labeled ddNTP. (click to expand)

Technical variations of chain-termination sequencing include tagging with nucleotides containing radioactive phosphorus for radiolabelling, or using a primer labeled at the 5’ end with a fluorescent dye. Dye-primer sequencing facilitates reading in an optical system for faster and more economical analysis and automation. The later development by Leroy Hood and coworkers [11][12] of fluorescently labeled ddNTPs and primers set the stage for automated, high-throughput DNA sequencing.

Sequence ladder by radioactive sequencing compared to fluorescent peaks (click to expand)

Chain-termination methods have greatly simplified DNA sequencing. For example, chain-termination-based kits are commercially available that contain the reagents needed for sequencing, pre-aliquoted and ready to use. Limitations include non-specific binding of the primer to the DNA, affecting accurate read-out of the DNA sequence, and DNA secondary structures affecting the fidelity of the sequence.

[edit]

Dye-terminator sequencing

Capillary electrophoresis (click to expand)

Dye-terminator sequencing utilizes labelling of the chain terminator ddNTPs, which permits sequencing in a single reaction, rather than four reactions as in the labelled-primer method. In dye-terminator sequencing, each of the four dideoxynucleotide chain terminators is labelled with fluorescent dyes, each of which with different wavelengths of fluorescence and emission. Owing to its greater expediency and speed, dye-terminator sequencing is now the mainstay in automated sequencing. Its limitations include dye effects due to differences in the incorporation of the dye-labelled chain terminators into the DNA fragment, resulting in unequal peak heights and shapes in the electronic DNA sequence trace chromatogram after capillary electrophoresis (see figure to the left). This problem has been addressed with the use of modified DNA polymerase enzyme systems and dyes that minimize incorporation variability, as well as methods for eliminating "dye blobs". The dye-terminator sequencing method, along with automated high-throughput DNA sequence analyzers, is now being used for the vast majority of sequencing projects.

[edit]

Challenges

Common challenges of DNA sequencing include poor quality in the first 15–40 bases of the sequence and deteriorating quality of sequencing traces after 700–900 bases. Base calling software typically gives an estimate of quality to aid in quality trimming.

In cases where DNA fragments are cloned before sequencing, the resulting sequence may contain parts of the cloning vector. In contrast, PCR-based cloning and emerging sequencing technologies based on pyrosequencing often avoid using cloning vectors. Recently, one-step Sanger sequencing (combined amplification and sequencing) methods such as Ampliseq and SeqSharp have been developed that allow rapid sequencing of target genes without cloning or prior amplification.[13] [14]

Current methods can directly sequence only relatively short (300–1000 nucleotides long) DNA fragments in a single reaction. The main obstacle to sequencing DNA fragments above this size limit is insufficient power of separation for resolving large DNA fragments that differ in length by only one nucleotide.

[edit]

Automation and sample preparation

View of the start of an example dye-terminator read (click to expand)

Automated DNA-sequencing instruments (DNA sequencers) can sequence up to 384 DNA samples in a single batch (run) 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.

[edit]

Amplification and clonal selection

Genomic DNA is fragmented into random pieces and cloned as a bacterial library. DNA from individual bacterial clones is sequenced and the sequence is assembled by using overlapping DNA regions.(click to expand)

Large-scale sequencing aims at sequencing very long DNA pieces, such as whole chromosomes. Common approaches consist of cutting (with restriction enzymes) or shearing (with mechanical forces) large DNA fragments into shorter DNA fragments. The fragmented DNA is cloned into a DNA vector, and amplified in Escherichia coli. Short DNA fragments purified from individual bacterial colonies are individually sequenced and assembled electronically into one long, contiguous sequence. This method does not require any pre-existing information about the sequence of the DNA and is referred to as de novo sequencing. Gaps in the assembled sequence may be filled by primer walking. The different strategies have different tradeoffs in speed and accuracy; shotgun methods are often used for sequencing large genomes, but its assembly is complex and difficult, particularly with sequence repeats often causing gaps in genome assembly.

Most sequencing approaches use an in vitro cloning step to amplify individual DNA molecules, because their molecular detection methods are not sensitive enough for single molecule sequencing. Emulsion PCR[15] isolates individual DNA molecules along with primer-coated beads in aqueous droplets within an oil phase. Polymerase chain reaction (PCR) then coats each bead with clonal copies of the DNA molecule followed by immobilization for later sequencing. Emulsion PCR is used in the methods by Marguilis et al. (commercialized by 454 Life Sciences), Shendure and Porreca et al. (also known as "Polony sequencing") and SOLiD sequencing, (developed by Agencourt, now Applied Biosystems).[16][17][18] Another method for in vitro clonal amplification is bridge PCR, where fragments are amplified upon primers attached to a solid surface, used in the Illumina Genome Analyzer. The single-molecule method developed by Stephen Quake's laboratory (later commercialized by Helicos) is an exception: it uses bright fluorophores and laser excitation to detect pyrosequencing events from individual DNA molecules fixed to a surface, eliminating the need for molecular amplification.[19]

[edit]

High-throughput sequencing

The high demand for low-cost sequencing has driven the development of high-throughput sequencing technologies that parallelize the sequencing process, producing thousands or millions of sequences at once.[20][21] High-throughput sequencing technologies are intended to lower the cost of DNA sequencing beyond what is possible with standard dye-terminator methods.[22]

[edit]

454 pyrosequencing

Main article: 454 Life Sciences#Technology

A parallelized version of pyrosequencing was developed by 454 Life Sciences. The method amplifies DNA inside water droplets in an oil solution (emulsion PCR), with each droplet containing a single DNA template attached to a single primer-coated bead that then forms a clonal colony. The sequencing machine contains many picolitre-volume wells each containing a single bead and sequencing enzymes. Pyrosequencing uses luciferase to generate light for detection of the individual nucleotides added to the nascent DNA, and the combined data are used to generate sequence read-outs.[16] This technology provides intermediate read length and price per base compared to Sanger sequencing on one end and Solexa and SOLiD on the other.[23]

[edit]

Solexa sequencing

Solexa has developed a sequencing technology based on reversible dye-terminators. DNA molecules are first attached to primers on a slide and amplified so that local clonal colonies are formed (bridge amplification). One type of nucleotide at a time is then added, and non-incorporated nucleotides are washed away. Unlike pyrosequencing, the DNA can only be extended one nucleotide at a time. A camera takes images of the fluorescently labeled nucleotides and the dye is chemically removed from the DNA, allowing a next cycle.[24]

[edit]

SOLiD sequencing

Main article: ABI Solid Sequencing

Applied Biosystems' SOLiD technology employs sequencing by ligation. Here, a pool of all possible oligonucleotides of a fixed length are labeled according to the sequenced position. Oligonucleotides are annealed and ligated; the preferential ligation by DNA ligase for matching sequences results in a signal informative of the nucleotide at that position. Before sequencing, the DNA is amplified by emulsion PCR. The resulting bead, each containing only copies of the same DNA molecule, are deposited on a glass slide.[25] Similar to Solexa sequencing, this technology produces short read lengths at a low price per base.[23]

[edit]

Future methods

Sequencing by hybridization is a non-enzymatic method that uses a DNA microarray. A single pool of DNA whose sequence is to be determined is fluorescently labeled and hybridized to an array containing known sequences. Strong hybridization signals from a given spot on the array identifies its sequence in the DNA being sequenced.[26] Mass spectrometry may be used to determine mass differences between DNA fragments produced in chain-termination reactions.[27]

DNA sequencing methods currently under development include labeling the DNA polymerase,[28] reading the sequence as a DNA strand transits through nanopores,[29][30] and microscopy-based techniques, such as AFM or electron microscopy that are used to identify the positions of individual nucleotides within long DNA fragments (>5,000 bp) by nucleotide labeling with heavier elements (e.g., halogens) for visual detection and recording.[31][32]

In microfluidic Sanger sequencing the entire thermocycling amplification of DNA fragments as well as their separation by electrophoresis is done on a single glass wafer (approximately 10 cm in diameter) thus reducing the reagent usage as well as cost.[citation needed] In some instances researchers[who?] have shown that they can increase the throughput of conventional sequencing through the use of microchips.[citation needed] Research will still need to be done in order to make this use of technology effective.

In October 2006, the X Prize Foundation established an initiative to promote the development of full genome sequencing technologies, called the Archon X Prize, intending to award $10 million to "the first Team that can build a device and use it to sequence 100 human genomes within 10 days or less, with an accuracy of no more than one error in every 100,000 bases sequenced, with sequences accurately covering at least 98% of the genome, and at a recurring cost of no more than $10,000 (US) per genome."[33]

DNA Sequencing

DNA sequencing is the determination of the precise sequence of nucleotides in a sample of DNA.

The most popular method for doing this is called the dideoxy method or Sanger method (named after its inventor, Frederick Sanger, who was awarded the 1980 Nobel prize in chemistry [his second] for this achievment).

DNA is synthesized from four deoxynucleotide triphosphates. The top formula shows one of them: deoxythymidine triphosphate (dTTP). Each new nucleotide is added to the 3′ -OH group of the last nucleotide added. Link to discussion of DNA synthesis.

The dideoxy method gets its name from the critical role played by synthetic nucleotides that lack the -OH at the 3′ carbon atom (red arrow). A dideoxynucleotide (dideoxythymidine triphosphate — ddTTP — is the one shown here) can be added to the growing DNA strand but when it is, chain elongation stops because there is no 3′ -OH for the next nucleotide to be attached to. For this reason, the dideoxy method is also called the chain termination method.

The bottom formula shows the structure of azidothymidine (AZT), a drug used to treat AIDS. AZT (which is also called zidovudine) is taken up by cells where it is converted into the triphosphate. The reverse transcriptase of the human immunodeficiency virus (HIV) prefers AZT triphosphate to the normal nucleotide (dTTP). Because AZT has no 3′ -OH group, DNA synthesis by reverse transcriptase halts when AZT triphosphate is incorporated in the growing DNA strand. Fortunately, the DNA polymerases of the host cell prefer dTTP, so side effects from the drug are not so severe as might have been predicted.

The Procedure

The DNA to be sequenced is prepared as a single strand.

This template DNA is supplied with

a mixture of all four normal (deoxy) nucleotides in ample quantities

dATP

dGTP

dCTP

dTTP

a mixture of all four dideoxynucleotides, each present in limiting quantities and each labeled with a "tag" that fluoresces a different color:

ddATP

ddGTP

ddCTP

ddTTP

DNA polymerase I

Because all four normal nucleotides are present, chain elongation proceeds normally until, by chance, DNA polymerase inserts a dideoxy nucleotide (shown as colored letters) instead of the normal deoxynucleotide (shown as vertical lines). If the ratio of normal nucleotide to the dideoxy versions is high enough, some DNA strands will succeed in adding several hundred nucleotides before insertion of the dideoxy version halts the process.

At the end of the incubation period, the fragments are separated by length from longest to shortest. The resolution is so good that a difference of one nucleotide is enough to separate that strand from the next shorter and next longer strand. Each of the four dideoxynucleotides fluoresces a different color when illuminated by a laser beam and an automatic scanner provides a printout of the sequence.