Sunday, 27 October 2019

DNA Techniques

DNA Techniques


"DNA Gel Electrophoresis"

DNA Gel Electrophoresis

The most common method used in DNA analysis is agarose gel electrophoresis of DNA. Agarose is a polysaccharide that forms a matrix, or gel, with nonuniform and relatively large pore- size gels, suitable for separation of DNA fragments of over 100 base pairs (for shorter fragments, other gel material, such as acrylamide, is commonly used). The negatively charged DNA molecule is thus “loaded” onto the gel, and is subjected to an electrical field, where the molecule moves toward the positively charged cathode. The shorter the DNA, the easier it is to pass through the agarose gel; hence, size separation of DNA molecules is possible.

Although not discussed in the main text, it should be noted that capillary electrophoresis has become a common microfluidic tool used in much contemporary equipment, particularly for automation and high- throughput assays. The system is mainly composed of sample vials, a capillary tube filled with an electrolyte, and a power supply. When voltage is applied to the sample in the capillary tube, the sample solution moves into the capillary by a combination of capillary action, pressure, and electrical field, and “moves” toward the cathode. The analytes within the sample will migrate differently within the capillary, analyzed with the help of a computer at the end of the capillary, even when the sample size is very small (and not suitable for conventional agarose electrophoresis). This system gives capillary electrophoresis another huge advantage over agarose gel electrophoresis. Multiple capillaries can run in parallel, generating a system suitable for high- throughput analysis.

Nucleic Acid Blotting

Nucleic Acid Blotting

Historically, perhaps the oldest blotting method that we know today was created by Edwin Southern, which he had devised so as to locate specific DNA sequences on the gel. The Southern blotting method essentially involves the digestion of the (genomic, amplified, etc.) DNA with a restriction enzyme, giving a mixture of DNA fragments of different sizes, which produces a “smear” when running on agarose gel electrophoresis. The DNA thus separated in the gel is then denatured by an alkaline treatment so that the DNA can be dissociated into single strands that can then be hybridized to the probes. The DNA is then transferred from the agarose gel to a (nylon) membrane and cross-linked, that is, immobilized. This membrane is then treated with a (usually radiolabeled) probe, which recognizes and hybridizes with homologous sequences. The membrane is washed to get rid of the excess probe and exposed to film or a phosphorimager cassette, after which the bands are visualized at the sites of hybridization.

Nucleic Acid Blotting

Figure. A schematic depiction of Southern blotting. The DNA samples are loaded into the wells, and then blotted onto a membrane. The membrane is then hybridized with a labeled probe against the target DNA. After autoradiography or other methods of visualization, any DNA that hybridizes to the probe is seen as a band on the membrane.

"Polymerase Chain Reaction"

Polymerase Chain Reaction

A polymerase chain reaction (PCR), developed by Kary Mullis in 1983, has been a routinely used method in almost every molecular biology laboratory for quite some time. Kary Mullis and Michael Smith were later awarded the Nobel Prize in Chemistry for this invention (10 years later in 1993). PCR entails amplifying a particular target DNA sequence in vitro, using heat-stable DNA polymerases. (Originally, Taq DNA polymerase from Thermus Aquaticus was used but various other enzymes have been isolated from different bacteria and exploited commercially since then.) In order to amplify a target sequence, one must have at least an idea about the sequence, since primers (short oligonucleotides bordering the target sequence) must be designed.

Polymerase Chain Reaction

Figure. A schematic depiction of PCR. 
(a) The DNA template, DNA polymerase, dNTPs, primers, and buffer are mixed in a reaction
tube. In Step 1, template DNA in the tube is denatured by heat; in Step 2, the temperature is lowered so the primers can anneal to their specific targets; in Step 3, DNA polymerase in the tube elongates from the primers and amplifies the DNA. This cycle is repeated around 30 times, yielding millions of copies of the target DNA region. 
(b) A gross simplification of primer design: primers are designed complementary to the borders of the target region, therefore the target sequence must be known (see specialty books or manuals for the details for designing primers). 
(c) If the exact sequence is not known (e.g., the protein sequence is known but the coding DNA sequence is not), or if the primer is designed to amplify the same DNA region from a number of different species (such as a human, mouse, and rat in the example, where some single nucleotide differences are possible), then a degenerate primer is to be designed. (Please note that the sequences and primers here are purely hypothetical, and similarly, the hypothetical primer designs shown are not optimal.)

These primers need to be as specific as possible for the target sequence; however, if, for example, only the protein sequence is known, then a coding sequence will be “predicted” using the genetic code table. Since multiple codons may code for the same amino acids, there will be many alternatives for the predicted coding sequence, and this will be reflected in the designing of so-called degenerate primers. Primers are typically around 18–20 bases in length, seldom going up to 30 bases, but they should have around 40 to 60% G- C content and have matching annealing temperatures. These temperatures are estimated as roughly 10°C lower than the melting temperature, the simplest and quickest formula of which is Tm = 2 × (n A + n T ) + 4 × (n G + n C )°C, where n is the number of the base indicated as subscript in the primer sequence. It is also a good idea to have one or two G or C residues at the end of the primer so that 3′ ends bind strongly to the template, necessary for efficient elongation. 

Once the primers are designed and a template is obtained, they will be incubated along with a DNA polymerase of choice, buffer, and deoxyribonucleotides, or dNTPs. In Step 1, template DNA will be denatured at a high temperature (usually 95°C), followed by Step 2, primer annealing. After the primers are allowed to anneal, DNA polymerase is set to elongate from the primers. This cycle is repeated around 30 times, generating millions of copies of the amplified region. These amplified regions can be visualized on an agarose gel, and the resulting DNA band can either be used for cloning, or sent for sequencing, and so forth, after purification.

"Real-Time PCR"

Real-Time PCR

A more recent modification of the PCR amplification is the so-called real-time PCR, or quantitative PCR (qPCR) since it employs fluorescent dyes to monitor the amplification process in real-time. There are many different versions of qPCR, either using a fluorescent DNA dye such as SYBR Green, which binds to double-stranded DNA (i.e., it is nonspecific), or using gene-specific probes that are fluorescently labeled. The principle behind the first strategy for qPCR is that since the fluorescent dye binds to double-stranded DNA, as the amount of DNA doubles up at each cycle (2 n ), the amount of fluorescence is doubled, which can be measured automatically at the end of each cycle (hence, real-time) and recorded. In the second strategy, since gene-specific probes will only bind to the target sequence, it is highly specific to the region that is amplified; the fluorescently labeled probe will bind to the amplified DNA at each cycle, increasing in intensity as the cycles progress, and as above, the fluorescence intensity is measured at the end of each cycle (again, real-time). The slight difference here is that normally the probe has both a fluorescent dye and a quencher, preventing any fluorescence emission from the unbound probe. When the annealing step is reached, both the probe and the primers bind to the target sequence, and during elongation, the DNA polymerase that extends the sequence from the primer will reach the probe, which leads to the degradation of the 5′-quencher due to the 5′-3′ exonuclease activity of the polymerase. The release of the quencher from the probe during this elongation step will free the fluorescent dye of the probe, and emission will be detected and recorded. Other than these details, the qPCR reaction is pretty much the same as a normal PCR reaction. It should be noted that there are many different variations of this original probe-based qPCR strategy that we will not discuss here in detail.

"DNA Sequencing"

DNA Sequencing

The DNA sequencing was initially developed in the 1970s, and the original method that became routinely used for sequencing DNA was developed by Maxam and Gilbert (i.e., known as Maxam– Gilbert sequencing) and is based on chemical modification of DNA, followed by site-specific cleavage of this modified DNA. Since this method relies on the use of hazardous chemicals and radioactivity, it is hardly ever used on a routine basis in molecular biology laboratories; therefore, we will not go into the details of this outdated method. Instead, we will introduce the more recent, dideoxynucleotide- based Sanger method, or the chain- termination method, which is also the basic principle in many of the modern automated sequencing technologies.

In Sanger or dideoxy sequencing (also known as the chain termination method), dideoxynucleotides (ddNTPs) are used in addition to the normal deoxynucleotides (dNTPs): since dideoxynucleotides lack both 2′- and 3′-OH groups, 5′-to-3′ extension between 5′-phosphate and 3′-OH groups cannot take place, hence “terminating” the elongation of new DNA synthesis. Essentially, DNA synthesis takes place from a single-stranded template using an oligonucleotide primer (called the sequencing primer) and dNTPs. After a brief period of incubation, this reaction is aliquotted into four different tubes, each of which contains a different labeled (radioactively or fluorescently), dideoxynucleotide terminator. Fluorescently labeled ddNTPs are more commonly used in automated sequencers, where different fluorescence emissions are observed by a detector, base by base, giving the sequence of the template.

"Next-Generation Sequencing"

The idea behind high- throughput technologies, in general, is to screen a large amount of data for a minimum price and in as short a time as possible. As such, it is not surprising that next-generation sequencers generate data that is equivalent to several hundred Sanger- type sequencers, operated by one person in one day (Mardis 2007; Schuster 2008). Recently, Roche Diagnostics released 454 pyrosequencing (also known as sequencing by synthesis) technologies, where DNA is captured by immobilization on beads and amplified inside water-oil emulsion. The machine uses many wells with picoliter volume to amplify from this template DNA, and uses luciferase for monitoring (the addition of new nucleotides results in a light signal); however, the cost per base that is sequenced is too high. Illumina, on the other hand, has developed another high- throughput method that uses reversible dye terminators (Metzker 2010).

Next-Generation Sequencing

As these nucleotides are added to the DNA one at a time, the fluorescence of the nucleotides is recorded. Last, ABI’s SOLiD (Sequencing by Oligonucleotide Ligation and Detection) sequencing relies on ligation, where oligonucleotides of a fixed length are captured on magnetic beads (a library of one fragment per bead), with an adapter sequence. When a universal primer anneals, a library of probes is added to the reaction, allowing the ligation of the probe to the primer, followed by ligation at each cycle (Mardis 2007).

Next-Generation Sequencing

Figure. A diagram of the dideoxy sequencing strategy. Unlabeled deoxynucleotides are used to briefly initiate the sequencing reaction, by DNA polymerase extending the nascent DNA with the help of a sequencing primer and dNTPs after denaturation of the template DNA. Thereafter, the reaction mix is typically aliquoted into four tubes, to each of which one dideoxynucleotide (ddNTP) is added. In each reaction tube, the DNA polymerase will incorporate normal dNTPs, but will also have an equal opportunity of incorporating a ddNTP. For example, in the ddCTP tube, the polymerase will either incorporate a dCTP or a ddCTP with equal probability opposite a “G” on the template strand. The resulting reaction products (which contain a mixture of different sized products in each tube) can either be run on a sequencing gel in classical Sanger dideoxy sequencing (where the nascent DNA will have to be radioactively labeled in the first part of the reaction), or in automated sequencing where the ddNTPs can each be labeled with a different fluorescent label (typically ddCTP is labeled blue, dTTP is labeled red, dATP is labeled with green, and ddGTP is labeled with yellow fluorescent dye), run on capillary electrophoresis and analyzed by computer.

The term deep sequencing is used in a different context— it refers to the number of times a sequence is read or the coverage. Deep sequencing is used for reads of seven times or more, whereas ultra-deep is now being used for over 100 reads, giving a higher coverage. This coverage is particularly useful if single nucleotide polymorphisms (SNP) are being investigated since in a single read there are likely to be sequencing errors, whereas when multiple reads are used, it becomes statistically more significant to determine which alterations are simply sequencing errors, and which ones are true SNPs.

Another breakthrough came with the application of microfluidics and microwell technologies to sequencing, making single-cell genomic DNA sequencing as well as RNA sequencing and transcriptomics possible, which was chosen as the “Method of the Year 2013” by Nature Methods (Editorial 2014b). This technology has the advantage of making clinical applications or work with rare or a limited number of cells far easier than any other technology.

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