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Substantial advances have been made in gene analysis and genomics in recent years, and this has been accelerated by the continued development and refinement of methods and techniques for studying nucleic acids. The application of molecular biology techniques has allowed understanding of cellular processes, both in normal and disease states. The advent of this type of DNA analysis has provided insight into the genetic make up of patients and their disease susceptibility and diagnostics. Prognostic analysis has also allowed the development of personalised or precision medicine and there is now great promise in further developments in drug discovery and molecular gene therapy. This chapter provides an overview of the general features of nucleic acid structure and function. It also describes some of the basic methods used in nucleic acid isolation and analysis, including restriction analysis, blotting, hybridization, the polymerase chain reaction (PCR) and associated methods, such as quantitative PCR and further genetic tests based on this method.

Major advances have been made in gene analysis and genomics in recent years and this has been accelerated by the continued development and refinement of methods and techniques for studying nucleic acids. One major area of current research is the identification and diagnosis of diseases that are multifactorial in nature. There are numerous diseases where analysis of genomes has provided insights into the disease and one particularly notable example is oncology. Molecular genetic analysis of this area has allowed a discrete set of cellular genes, termed oncogenes and tumour suppressor genes, to be identified and characterised. These genes and the proteins and enzymes they encode are major components of cell signalling and the cell cycle and are intimately involved in many aspects cell regulation. The disruption of oncogenes and tumour suppressor genes contributes to the early changes required for cancers to develop. Identification and analysis of these genes and genomes has already provided information for use in diagnostics and prognostics and a number have been shown to be biomarkers of a particular cancer type. In a number of cancers well-defined molecular events have been correlated with mutations in oncogenes and therefore in the corresponding protein. It is already possible to screen and predict the outcome of some disease processes at an early stage, a point which itself raises significant ethical dilemmas. The application of molecular biology has allowed understanding of cellular processes both in normal and disease states. The advent of this type of analysis has given rise to the development of personalised or precision medicine and there is now great promise in further developments in drug discovery and molecular gene therapy. A number of genetically engineered therapeutic proteins and enzymes have been developed and are already having an effect on disease management. In addition the correction of disorders at the gene level using gene therapy is also under way. Perhaps one of the most startling applications of molecular biology to date is indeed gene editing and the development of gene modifications methods such as the clustered regularly interspaced short palindromic repeats (CRISPR)–CRISPR-associated system 9 (cas9) system, which may have a profound effect on treating genetic-based diseases. In considering the potential utility of molecular biology techniques it is important to understand the basic structure of nucleic acids and gain an appreciation of how this dictates the function in vivo and in vitro. Indeed, many techniques used in molecular biology mimic in some way the natural functions of nucleic acids, such as replication and transcription. This chapter is intended to provide an overview of the general features of nucleic acid structure and function and describe some of the basic methods used in their isolation and analysis.

DNA and RNA are macromolecular structures composed of regular repeating polymers formed from nucleotides.1  These are the basic building blocks of nucleic acids and are derived from nucleosides, which are composed of two elements: a five-membered pentose carbon sugar (2-deoxyribose in DNA and ribose in RNA), and a nitrogenous base (Figure 1.1). The carbon atoms of the sugar are designated ‘prime’ (l′, 2′, 3′, etc.). To distinguish them from the carbons of the nitrogenous bases, of which there are two types, either a purine or a pyrimidine. A nucleotide, or nucleoside phosphate, is formed by the attachment of a phosphate to the 5′ position of a nucleoside by an ester linkage. Such nucleotides can be joined together by the formation of a second ester bond by reaction between the phosphate of one nucleotide and the 3′ hydroxyl of another, thus generating a 5′ to 3′ phosphodiester bond between adjacent sugars; this process can be repeated indefinitely to give long polynucleotide molecules. DNA has two such polynucleotide strands. However, since each strand has both a free 5′ hydroxyl group at one end, and a free 3′ hydroxyl at the other end, each strand has a polarity or directionality. The polarities of the two strands of the molecule are in opposite directions, and thus DNA is described as an ‘anti-parallel’ structure.

Figure 1.1

Representation of a deoxynucleoside triphosphate indicating the three components of a sugar, triphosphate and a base. The base can be either A C G or T. In RNA the 2′ carbon has an OH whereas it is deoxy in DNA. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.1

Representation of a deoxynucleoside triphosphate indicating the three components of a sugar, triphosphate and a base. The base can be either A C G or T. In RNA the 2′ carbon has an OH whereas it is deoxy in DNA. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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The purine bases (composed of fused five- and six-membered rings), adenine (A) and guanine (G), are found in both RNA and DNA, as is the pyrimidine (a single six-membered ring) cytosine (C). The other pyrimidines are each restricted to one type of nucleic acid: uracil (U) occurs exclusively in RNA, whilst thymine (T) is limited to DNA. Thus it is possible to distinguish between RNA and DNA on the basis of the presence of ribose and uracil in RNA, and deoxyribose and thymine in DNA. However, it is the sequence of bases along the structure, which distinguishes one DNA (or RNA) from another.

The two polynucleotide chains in DNA are usually found in the shape of a right-handed double helix, in which the bases of the two strands lie in the centre of the molecule, with the sugar–phosphate backbones on the outside.2  A crucial feature of this double-stranded structure is that it depends on the sequence of bases in one strand being complementary to those in the other strand. A purine base attached to a sugar residue on one strand is always hydrogen bonded to a pyrimidine base attached to a sugar residue on the other strand. Moreover, adenine (A) always pairs with thymine (T) or uracil (U) in RNA, via two hydrogen bonds, and guanine (G) always pairs with cytosine (C) by three hydrogen bonds (Figure 1.2). When these conditions are met a stable double-helical structure results in which the backbones of the two strands are, on average, a constant distance apart. Thus, if the sequence of one strand is known, that of the other strand can be deduced. The strands are designated as plus (+) and minus (−) and an RNA molecule complementary to the minus (−) strand is synthesised during transcription. The base sequence may cause significant local variations in the shape of the DNA molecule, and these variations are vital for specific interactions between the DNA and various proteins to take place. Although the three-dimensional structure of DNA may vary it generally adopts a double helical structure termed the b form or b-DNA in vivo.

Figure 1.2

Representation of the four bases in DNA and their complementary base pairing, A–T and C–G through hydrogen bonds. The right hand side depicts a DNA double helix. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.2

Representation of the four bases in DNA and their complementary base pairing, A–T and C–G through hydrogen bonds. The right hand side depicts a DNA double helix. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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It has been well recognized for some time that DNA as a structure may be chemically modified without the underlying DNA sequence being altered. One of the most important modifications is the addition of a methyl (CH3) group to cytosine, termed DNA methylation and catalysed by DNA methyltransferases. This results in what some describe as the fifth base in DNA. Approximately 1.5% of human DNA is methylated (termed the epigenome) and the methylation status appears to have a profound effect on gene expression, where hypomethylation appears to promote gene expression. This feature of gene expression control is termed epigenetics and is a complex process which is also extended to the modification of histone proteins and some small RNA molecules involved in gene expression control. Importantly, epigenetics appears to play a role in disease states, such as cancers and certain neurological diseases, which may lead to a new means of future treatment.

The two anti-parallel strands of DNA are held together only by the weak forces of hydrogen bonding between complementary bases, and partly by hydrophobic interactions between adjacent, stacked base pairs, termed base-stacking. Little energy is needed to separate a few base pairs, and so, at any instant, a few short stretches of DNA will be opened up to the single-stranded conformation. However, such stretches immediately pair up again at room temperature, so the molecule as a whole remains predominantly double-stranded.

If, however, a DNA solution is heated to approximately 90 °C or above there will be enough kinetic energy to denature the DNA completely, causing it to separate into single strands. The temperature at which 50% of the DNA is melted is termed the melting temperature or Tm, and this depends on the nature of the DNA. If several different samples of DNA are melted, it is found that the Tm is highest for those DNA molecules that contain the highest proportion of cytosine and guanine, and Tm can actually be used to estimate the percentage (C + G) in a DNA sample. This relationship between Tm and (C + G) content arises because cytosine and guanine form three hydrogen bonds when base-paired, whereas thymine and adenine form only two. Because of the differential numbers of hydrogen bonds between A–T and C–G pairs those sequences with a predominance of C–G pairs will require greater energy to separate or denature them. The conditions required to separate a particular nucleotide sequence are also dependent on environmental conditions, such as salt concentration. If denatured DNA is cooled it is possible for the separated strands to re-associate, a process known as renaturation.

Strands of RNA and DNA will associate with each other, if their sequences are complementary, to give double-stranded, hybrid molecules. Similarly, strands of labelled RNA or DNA, when added to a denatured DNA preparation, will act as probes for DNA molecules to which they are complementary. This hybridisation of complementary strands of nucleic acids is a cornerstone for many molecular biology techniques and is very useful for isolating a specific fragment of DNA from a complex mixture. It is also possible for small single-stranded fragments of DNA (up to 40 bases in length), termed oligonucleotides, to hybridise to a denatured sample of DNA. This type of hybridisation is termed annealing and again is dependent on the base sequence of the oligonucleotide and the salt concentration of the sample.

The use of DNA for analysis or manipulation usually requires that it is isolated and purified to a certain extent.3  DNA is recovered from cells by the gentlest possible method of cell rupture to prevent the DNA from fragmenting by mechanical shearing. This is usually in the presence of ethylenediaminetetraacetic acid (EDTA) which chelates the Mg2+ ions needed for enzymes that degrade DNA, termed DNAse. Ideally, cell walls, if present, should be digested enzymatically (e.g. by lysozyme treatment of bacteria), and the cell membrane should be solubilised using detergent. If physical disruption is necessary it should be kept to a minimum and should involve cutting or squashing of cells, rather than the use of shear forces. Cell disruption (and most subsequent steps) should be performed at 4 °C, using glassware and solutions which have been autoclaved to destroy DNAse activity (Figure 1.3).

Figure 1.3

General steps involved in extracting DNA from cells or tissues. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.3

General steps involved in extracting DNA from cells or tissues. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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After release of nucleic acids from the cells, RNA can be removed by treatment with ribonuclease (RNAse) that has been heat-treated to inactivate any DNAse contaminants; RNAse is relatively stable to heat as a result of its disulphide bonds, which ensure rapid renaturation of the molecule on cooling. The other major contaminant, protein, is removed by shaking the solution gently with water-saturated phenol, or with a phenol–chloroform mixture, either of which will denature proteins but not nucleic acids. Centrifugation of the emulsion formed by this mixing produces a lower, organic phase, separated from the upper, aqueous phase by an interface of denatured protein. The aqueous solution is recovered and deproteinised repeatedly, until no more material is seen at the interface. Finally, the deproteinised DNA preparation is mixed with two volumes of absolute ethanol and the DNA is allowed to precipitate out of solution in a freezer. After centrifugation, the DNA pellet is redissolved in a buffer containing EDTA to inactivate any DNAses present. This solution can be stored at 4 °C for at least a month. DNA solutions can be stored frozen, although repeated freezing and thawing tends to damage long DNA molecules by shearing.

The procedure described is suitable for total cellular DNA. If the DNA from a specific organelle or viral particle is needed, it is best to isolate the organelle or virus before extracting its DNA, since the recovery of a particular type of DNA from a mixture is usually rather difficult. Where a high degree of purity is required DNA may be subjected to density gradient ultracentrifugation through caesium chloride which is particularly useful for the preparation plasmid DNA. It is possible to check the integrity of the DNA by agarose gel electrophoresis and determine the concentration of the DNA by using the fact that 1 absorbance unit at a wavelength of 260 nm (A260) equates to 50 µg ml−1 of DNA and thus:

Contaminants may also be identified in the sample by employing scanning UV-spectrophotometry from 200 nm to 300 nm. A ratio of A260 : A280 of approximately 1.8 indicates that the sample is free of protein contamination, which absorbs strongly at 280 nm.

The methods used for RNA isolation are very similar to those described above for DNA; however, RNA molecules are relatively short and therefore less easily damaged by shearing, so cell disruption can be rather more vigorous. RNA is, however, vulnerable to digestion by RNAses, which are present endogenously in various concentrations in certain cell types and exogenously on fingers. Gloves should therefore be worn, and a strong detergent should be included in the isolation medium to immediately denature any RNAses. Subsequent deproteinization should be particularly rigorous, since RNA is often tightly associated with proteins. DNAse treatment can be used to remove DNA, and RNA can be precipitated by ethanol. One reagent which is commonly used in RNA extraction is guanidinium thiocyanate (GTC) which is both a strong inhibitor of RNAse and a protein denaturant. It is possible to check the integrity of an RNA extract by analysing it by agarose gel electrophoresis. The most abundant RNA species are the rRNA molecules. For prokaryotes these are 16S and 23S and for eukaryotes the molecules are slightly heavier at 18S and 28S. These appear as discrete bands following agarose gel electrophoresis and, importantly, if intact indicate that the other RNA components, such as mRNA, are likely to be intact also. Electrophoresis is usually carried out under denaturing conditions to prevent secondary structure formation in the RNA. The concentration of the RNA may be estimated by using UV-spectrophotometry. At 260 nm 1 absorbance unit equates to 40 µg ml−1 of RNA and therefore:

Contaminants may also be identified in the same way as for DNA by scanning UV-spectrophotometry, however in the case of RNA an A260 : A280 ratio of approximately 2 would be expected for a sample containing no contaminants.

In many cases it is desirable to isolate eukaryotic mRNA which constitutes only 2–5% of cellular RNA from a mixture of total RNA molecules. This may be carried out by affinity chromatography on oligo(dT)-cellulose columns. At high salt concentrations, the mRNA containing poly(A) tails binds to the complementary oligo(dT) molecules of the affinity column, and so mRNA will be retained; all other RNA molecules can be washed through the column by further high-salt solution. Finally, the bound mRNA can be eluted using a low concentration of salt. Nucleic acid species may also be subfractionated by more physical means such as electrophoretic or chromatographic separations based on differences in nucleic acid fragment sizes or physicochemical characteristics.

Many current molecular biology methods and their reagents can now be found in the form of a kit from a manufacturer, such as Sigma, or can be automated. The extraction of nucleic acids by these means is no exception. The advantage of their use lies in the fact that the reagents are standardised and quality control tested, providing a high degree of reliability. For example, glass bead preparations for DNA purification have been used increasingly and with reliable results. Small compact column-type preparations, such as Qiagen spin columns, are also used extensively in research and in routine DNA extraction. Essentially the same reagents for nucleic acid extraction may be used in a format that allows reliable and automated extraction. The process can also be automated with a low-throughput Qiacube system. Further methods are also available using kit-based extraction methods for RNA; these in particular have overcome some of the problems of RNA extraction, such as RNAse contamination. A number of fully automated nucleic acid extraction machines, such as the Qiasymphony system are now employed in areas where high throughput is required, e.g. clinical diagnostic laboratories. Here the raw samples, such as blood specimens, are placed in 96- or 384-well microtitre plates and these undergo a set computer-controlled processing pattern carried out robotically. Thus the samples are rapidly manipulated and extracted in approximately 45 min without any manual operations being undertaken. Nucleic acids can be extracted from a variety of samples, including blood, serum, frozen tissue sections, formalin-fixed paraffin-embedded (FFPE) tissue sections and biopsies among others, and all have their own unique challenges in the extraction process. Reliable and effective methods are a crucial element for further analysis, such as DNA sequencing and long-term storage in DNA banks.

A number of methods have been developed for the extraction of nucleic acids in single cells or completely cell-free.4  In addition to DNA, mRNA and microRNA (miRNA) and the various individual components have also been successfully isolated and assayed. Single-cell analysis relies on the preparation of single cells from a mixed population of cells and tissues. This can be achieved using a number of methods, although one of the most common is laser capture microdissection (LCM). Here, a few cells or single cells are typically prepared by a UV pulsed laser under automated control, or by manual microscopy of the tissue. The cells are removed from the surrounding tissue and captured for further analysis, such as nucleic acid amplification and DNA sequencing.

In addition to single-cell analysis cell-free DNA has been found in the circulation and has potential utility, especially in cases of fetal DNA analysis in the maternal bloodstream. This is proving useful as cell-free fetal DNA (cffDNA) is a very promising non-invasive method that can be used with various DNA analysis techniques to determine a number of genetic disorders. In a similar way circulating tumour cells (CTC) and circulating tumour DNA (ctDNA) may also be assayed for a number of diseases, including cancers. Of particular interest is the concept of the liquid biopsy using ctDNA. These are small fragments of DNA of approximately 170 base pairs (bp) originating from the tumour cells found in the bloodstream and which may be analysed using amplification methods, such as the polymerase chain reaction (PCR). This allows for the identification of biomarkers of the disease (e.g. KRAS) and mutations within them. However the collection of samples is not without problems, especially when standard methods of blood collection are used, e.g. heparin and when employing EDTA tubes. Furthermore white cell genomic DNA will be released from the samples if the methods are not optimised and this may interfere with the ctDNA analysis. Various methods of stabilisation have therefore been used to overcome these problems. The use of liquid biopsy is gaining interest as not only is it a non-invasive process but a number of studies have reported its ability to not only aid in diagnostics and screening but also to be of benefit in prognostics. It may further be an effective means of monitoring disease progression and response to treatment and a sensitive way of identifying minimal residual disease (MRD) found in reoccurrence of certain tumours.

The discovery and characterisation of a number of key enzymes has enabled the development of various techniques for the analysis and manipulation of DNA. In particular, the enzymes termed type II restriction endonucleases have come to play a key role in all aspects of molecular biology.5  These enzymes recognise certain DNA sequences, usually 4–6 bp in length, and cleave them in a defined manner. The sequences recognised are palindromic or of an inverted repeat nature (Figure 1.4). That is they read the same in both directions on each strand. When cleaved they leave a flush-ended or staggered (also termed a cohesive-ended) fragment depending on the particular enzyme used (Table 1.1). An important property of staggered ends is that those produced from different molecules by the same enzyme are complementary (or ‘sticky’) and so will anneal to each other. The annealed strands are held together only by hydrogen bonding between complementary bases on opposite strands. Covalent joining of the ends of each of the two strands may be carried out using the enzyme DNA ligase. This is widely exploited in molecular biology to enable the construction of recombinant DNA i.e. the joining of DNA fragments from different sources. Approximately 500 restriction enzymes have been characterised that recognise over 100 different target sequences. A number of these, termed isoschizomers, recognise different target sequences but produce the same staggered ends or overhangs. In addition restriction enzymes may be applied to various situations in molecular biology as indicated in Figure 1.5.

Figure 1.4

The cleavage of a DNA strand with a target site for the restriction enzyme EcoR1 indicating the ends of the DNA formed following digestion. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.4

The cleavage of a DNA strand with a target site for the restriction enzyme EcoR1 indicating the ends of the DNA formed following digestion. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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Table 1.1

Examples of restrictions enzymes with four-, six- or eight-base recognition sequences. Reproduced from ref. 22 with permission from the Royal Society of Chemistry

NameRecognition SequenceDigestion Products
Four-nucleotide recognition sequence  
 
HaeIII 5′-GGCC-3′ 5′-GG CC-3′ Blunt end digestion 
3′-CCGG-5′ 3′-CC GG-5′ 
HpaII 5′-CCGG-3′ 5′-C CGG-3′ Cohesive end digestion 
3′-GGCC-5′ 3′-GGC C-5′ 
Six-nucleotide recognition sequence  
 
BamHI 5′-GGATTC-3′ 5′-G GATCC-3′ 
3′-GGCC-5′ 3′-CCTAG G-5′ 
 
EcoRI 5′-GAATTC-3′ 5′-G AATCC-3′ 
3′-CTTAAG-5′ 3′-CTTAA G-5′ 
 
HindIII 5′-AAGCTT-3′ 5′-A AGCTT-3′ 
3′-TTCGAA-5′ 3′-TTCGA A-5′ 
Eight-nucleotide recognition sequence  
 
Not5′-GCGGCCGC-3′ 5′-GC GGCCGC-3′ 
3′-CGCCGGCG-5′ 3′-CGCCGG CG-5′ 
NameRecognition SequenceDigestion Products
Four-nucleotide recognition sequence  
 
HaeIII 5′-GGCC-3′ 5′-GG CC-3′ Blunt end digestion 
3′-CCGG-5′ 3′-CC GG-5′ 
HpaII 5′-CCGG-3′ 5′-C CGG-3′ Cohesive end digestion 
3′-GGCC-5′ 3′-GGC C-5′ 
Six-nucleotide recognition sequence  
 
BamHI 5′-GGATTC-3′ 5′-G GATCC-3′ 
3′-GGCC-5′ 3′-CCTAG G-5′ 
 
EcoRI 5′-GAATTC-3′ 5′-G AATCC-3′ 
3′-CTTAAG-5′ 3′-CTTAA G-5′ 
 
HindIII 5′-AAGCTT-3′ 5′-A AGCTT-3′ 
3′-TTCGAA-5′ 3′-TTCGA A-5′ 
Eight-nucleotide recognition sequence  
 
Not5′-GCGGCCGC-3′ 5′-GC GGCCGC-3′ 
3′-CGCCGGCG-5′ 3′-CGCCGG CG-5′ 
Figure 1.5

Indication of the application of restriction enzymes and the restriction fragment lengths generated in various situations. For example in A digestion with enzyme 1 and 2 results in a particular fragment size whereas in B a point mutation resulting in the abolition of enzyme 1 will result in a different fragment size. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.5

Indication of the application of restriction enzymes and the restriction fragment lengths generated in various situations. For example in A digestion with enzyme 1 and 2 results in a particular fragment size whereas in B a point mutation resulting in the abolition of enzyme 1 will result in a different fragment size. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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Electrophoresis in agarose or polyacrylamide gels is the most usual way to separate DNA molecules according to size. The technique can be used analytically or preparatively and can be qualitative or quantitative. Large fragments of DNA, such as chromosomes, may also be separated by a modification of electrophoresis termed pulsed field gel electrophoresis (PFGE). The easiest and most widely applicable method is electrophoresis in horizontal agarose gels, followed by staining with ethidium bromide. This dye binds to DNA by insertion between stacked base pairs (intercalation), and it exhibits a strong orange or red fluorescence when illuminated with ultraviolet light. Very often electrophoresis is used to check the purity and intactness of a DNA preparation or to assess the extent of an enzymatic reaction during, for example, the steps involved in the cloning of DNA (Figure 1.6). For such checks ‘mini-gels’ are particularly convenient, since they need little preparation, use small samples and provide results quickly. Agarose gels can be used to separate molecules larger than about 100 bp. For higher resolution or for the effective separation of shorter DNA molecules polyacrylamide gels are the preferred method.

Figure 1.6

A schematic illustration of a typical horizontal agarose gel electrophoresis set up for the separation of nucleic acids. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.6

A schematic illustration of a typical horizontal agarose gel electrophoresis set up for the separation of nucleic acids. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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When electrophoresis is used preparatively, the piece of gel containing the desired DNA fragment is physically removed with a scalpel. The DNA may be recovered from the gel fragment in various ways. This may include crushing with a glass rod in a small volume of buffer, using agarase to digest the agarose leaving the DNA, or by the process of electroelution. In this method the piece of gel is sealed in a length of dialysis tubing containing buffer and is then placed between two electrodes in a tank containing more buffer. Passage of an electrical current between the electrodes causes DNA to migrate out of the gel piece but it remains trapped within the dialysis tubing and can therefore be recovered easily.

Gel electrophoresis remains the established method for the separation and analysis of nucleic acids. However, a number of automated systems using pre-cast gels and standardised reagents are available that are now very popular. These are especially useful in situations where a large number of samples or high-throughput analysis is required. In addition, technologies, such as the Agilent bioanalyser, have been developed that obviate the need to prepare electrophoretic gels. These employ microfluidic circuits constructed on small cassette units that contain interconnected micro-reservoirs. The sample is applied to one area and driven through microchannels under computer-controlled electrophoresis. The channels lead to reservoirs allowing, for example, incubation with other reagents, such as dyes, for a specified time. Electrophoretic separation is thus carried out in a microscale format. The small sample size minimises sample and reagent consumption and the units, being computer controlled, allow data to be captured within a very short timescale. In addition dedicated spectrophotometers such as the Thermofisher Scientific Nanodrop system can provide nucleic acid concentrations quickly with limited sample volume. Alternative methods of analysis, including high-performance liquid chromatography-based approaches, have gained in popularity, especially for DNA mutation analysis. Mass spectrometry methods traditionally used in protein analysis, such as matrix-assisted laser desorption and ionization-time of flight (MALDI-TOF) are also becoming increasingly used for nucleic acid analysis due to their rapidity and increasing reliability.

Electrophoresis of DNA restriction fragments allows separation based on size to be carried out. However, it provides no indication as to the presence of a specific, desired fragment among the complex sample. This can be achieved by transferring the DNA from the intact gel onto a piece of nitrocellulose or nylon membrane placed in contact with it. This provides a more permanent record of the sample since DNA begins to diffuse out of a gel that is left for a few hours. First the gel is soaked in alkali to render the DNA single stranded. It is then transferred to the membrane so that the DNA becomes bound to it in exactly the same pattern as that originally on the gel. This transfer, named a Southern blot after its inventor Ed Southern, can be performed electrophoretically, or by drawing large volumes of buffer through both gel and membrane, thus transferring DNA from one to the other by capillary action.6  The point of this operation is that the membrane can now be treated with a labelled DNA molecule, for example a gene probe. This single-stranded DNA probe will hybridise under the right conditions to complementary fragments immobilised onto the membrane (Figure 1.7). The conditions of hybridisation, including the temperature and salt concentration, are critical for this process to take place effectively. This is usually referred to as the stringency of the hybridisation and it is particular to each individual gene probe and for each sample of DNA. A series of washing steps with buffer is then carried out to remove any unbound probe and the membrane is developed, after which the precise location of the probe and its target may be visualised. It is also possible to analyse DNA from different species or organisms by blotting the DNA and then using a gene probe representing a protein or enzyme from one of the organisms. In this way it is possible to search for related genes in different species. This technique is generally termed zoo blotting.

Figure 1.7

Steps involved in a Southern blot, note that the denaturation step can be achieved using sodium hydroxide before the blotting stage with a nylon membrane. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.7

Steps involved in a Southern blot, note that the denaturation step can be achieved using sodium hydroxide before the blotting stage with a nylon membrane. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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The same basic process of nucleic acid blotting can be used to transfer RNA from gels onto similar membranes.7  This allows the identification of specific mRNA sequences of a defined length by hybridisation to a labelled gene probe and is known as northern blotting. It is possible with this technique to not only detect specific mRNA molecules but also to quantify the relative amounts of the specific mRNA. It is usual to separate the mRNA transcripts by gel electrophoresis under denaturing conditions since this improves resolution and allows a more accurate estimation of the sizes of the transcripts. The format of the blotting may be altered from transfer from a gel to direct application to slots on a specific blotting apparatus containing the nylon membrane. This is termed slot or dot blotting and provides a convenient means of measuring the abundance of specific mRNA transcripts without the need for gel electrophoresis. It does not, however, provide information regarding the size of the fragments.

A further method of RNA analysis that overcomes the problems of RNA blotting is termed the ribonuclease protection assay (RPA). This is a solution-based method where a probe that is complementary to the mRNA of interest is bound to form a hybrid that is resistant to digestion with RNAse. Thus while other single-stranded RNA molecules are digested, the intact hybrids can be further analysed by gel electrophoresis.

The availability of a gene probe is essential in many molecular biology techniques yet in many cases is one of the most difficult steps.8  The information needed to produce a gene probe, or a primer as used in PCR, may come from many sources but with the development and sophistication of bioinformatics and genetic databases computer-based analysis is usually one of the first stages (see Chapter 5). In some cases, it is possible to use related proteins from the same gene family to gain information on the most useful DNA sequence. Similar proteins or DNA sequences but from different species may also provide a starting point with which to produce a so-called heterologous gene probe. Although in some cases probes are already produced and cloned it is possible, armed with a DNA sequence from a DNA database, to chemically synthesise a single-stranded oligonucleotide probe. This is usually undertaken by computer-controlled gene synthesisers, which link deoxynucleotide triphosphates (dNTPs) together based on a desired sequence. It is essential to carry out certain checks before probe production to determine that the probe is unique, is not able to self-anneal and that it is not self complementary, all of which may compromise its use (Figure 1.8).

Figure 1.8

Various methods and applications in gene probe and primer production. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.8

Various methods and applications in gene probe and primer production. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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Where little DNA information is available to prepare a gene probe it is possible, in some cases, to use the knowledge gained from analysis of the corresponding protein. Thus it is possible to isolate and purify proteins and sequence part of the N-terminal end of the protein. From our knowledge of the genetic code, it is possible to predict the various DNA sequences that could code for the protein, and then synthesise appropriate oligonucleotide sequences chemically. Due to the degeneracy of the genetic code most amino acids are coded for by more than one codon, therefore there will be more than one possible nucleotide sequence which could code for a given polypeptide. The longer the polypeptide, the greater the number of possible oligonucleotides which must be synthesised. Fortunately, there is no need to synthesise a sequence longer than about 20 bases, since this should hybridise efficiently with any complementary sequences and should be specific for one gene. Ideally, a section of the protein should be chosen which contains as many tryptophan and methionine residues as possible, since these have unique codons, and there will therefore be fewer possible base sequences which could code for that part of the protein. The synthetic oligonucleotides can then be used as probes in a number of molecular biology methods.

An essential feature of a gene probe (and possibly a primer used in PCR) is that it can be visualised by some means. In this way a gene probe that hybridises to a complementary sequence may be detected and used to identify that desired sequence from a complex mixture. There are two main ways of labelling gene probes. Fluorescent labelling is now a popular method for tagging nucleic acids and includes dyes such as fluorescein amidite (FAM) and digoxigenin-labelled nucleotides. Additionally, fluorescent dyes, such as 4′,6-diamidino-2-phenylindole (DAPI) Picogreen and Ribogreen, are commonly used. Radioactive labelling with 32 phosphorous (32P), or for certain techniques 35 sulphur (35S) and tritium (3H), can also be used. These may be detected by the process of autoradiography where the labelled probe molecule, bound to sample DNA, located, for example, on a nylon membrane, is placed in contact with an X-ray-sensitive film. Following exposure, the film is developed and fixed just as a black and white negative and reveals the precise location of the labelled probe and therefore the DNA to which it has hybridised.

Non-radioactive fluorescent labels are increasingly being used to label DNA gene probes and now many have similar sensitivities which, when combined with their improved safety, have led to their widespread use.

The labelling systems are termed either direct or indirect. Direct labelling allows an enzyme reporter, such as alkaline phosphatase, to be coupled directly to the DNA. Although this may alter the characteristics of the DNA gene probe it offers the advantage of rapid analysis since no intermediate steps are needed. However indirect labelling is, at present, more popular. This relies on the incorporation of a nucleotide that has a label attached. At present three of the main labels in use are biotin, fluorescein and digoxigenin. These molecules are covalently linked to nucleotides using a carbon spacer arm of 7, 14 or 21 atoms. Specific binding proteins may then be used as a bridge between the nucleotide and a reporter protein, such as an enzyme. For example, biotin incorporated into a DNA fragment is recognised with a very high affinity by the protein streptavidin. This may either be coupled or conjugated to a reporter enzyme molecule, such as alkaline phosphatase. This is able to convert a colourless substrate para nitrophenol phosphate (PNPP) into the yellow compound para nitrophenol (PNP) and also offers a means of signal amplification. Alternatively, labels, such as digoxigenin, incorporated into DNA sequences may be detected by monoclonal antibodies, again conjugated to reporter molecules, including alkaline phosphatase. Thus, rather than the detection system relying on autoradiography which is necessary for radiolabels, a series of reactions resulting in either a colour, light or a chemiluminescence reaction takes place. This has important practical implications since autoradiography may take one to three days whereas colour and chemiluminescence reactions take minutes.

The simplest form of labelling DNA is by 5′ or 3′ end-labelling. 5′ end-labelling involves a phosphate transfer or exchange reaction where the 5′ phosphate of the DNA to be used as the probe is removed and in its place a labelled phosphate, traditionally 32P, is added. This is usually carried out by using two enzymes, the first, alkaline phosphatase, is used to remove the existing phosphate group from the DNA. Following removal of the released phosphate from the DNA a second enzyme, polynucleotide kinase, is added which catalyses the transfer of a phosphate group (32P labelled) to the 5′ end of the DNA. The newly labelled probe is then purified, usually by chromatography through a sephadex column, and may be used directly (Figure 1.9).

Figure 1.9

End-labelling of a gene probe at the 5′ end with alkaline phosphatase and polynucleotide kinase. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.9

End-labelling of a gene probe at the 5′ end with alkaline phosphatase and polynucleotide kinase. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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Using the other end of the DNA molecule, the 3′ end, is slightly less complex. Here a new dNTP which is labelled (e.g.32PαdATP or biotin-labelled dNTP) is added to the 3′ end of the DNA by the enzyme terminal transferase. Although this is a simpler reaction a potential problem exists because a new nucleotide is added to the existing sequence and so the complete sequence of the DNA is altered, which may affect its hybridisation to its target sequence. End-labelling methods also suffer from the fact that only add one label is added to the DNA, so they are of a lower activity in comparison with methods that incorporate label along the length of the DNA. Fluorescent labels, such as FAM, are now used as an alternative to radiolabels and further developments in probe technology, such as peptide nucleic acid (PNA), locked nucleic acid (LNA) or minor groove binding (MGB) probes, are also widely available.

The DNA to be labelled is first denatured and then placed under renaturing conditions in the presence of a mixture of many different random sequences of hexamers or hexanucleotides. These hexamers will, by chance, bind to the DNA sample wherever they encounter a complementary sequence and so the DNA will rapidly acquire an approximately random sprinkling of hexanucleotides annealed to it. Each of the hexamers can act as a primer for the synthesis of a fresh strand of DNA catalysed by DNA polymerase since it has an exposed 3′ hydroxyl group. The Klenow fragment of DNA polymerase is used for random primer labelling because it lacks a 5′–3′ exonuclease activity. This is prepared by cleavage of DNA polymerase with subtilisin, giving a large enzyme fragment which has no 5′–3′ exonuclease activity but which still acts as a 5′–3′ polymerase. Thus, when the Klenow enzyme is mixed with the annealed DNA sample in the presence of dNTPs, including at least one which is labelled, many short stretches of labelled DNA will be generated. In a similar way to random primer labelling the polymerase chain reaction may also be used to incorporate radioactive or non radioactive labels (Figure 1.10).

Figure 1.10

Random primer gene probe labelling. Random primers are incorporated and used as start points for DNA polymerase to synthesise a complementary strand of DNA whilst incorporating a labelled dNTP at complementary sites. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.10

Random primer gene probe labelling. Random primers are incorporated and used as start points for DNA polymerase to synthesise a complementary strand of DNA whilst incorporating a labelled dNTP at complementary sites. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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There have been a number of key developments in molecular biology, however, one that has had the most impact in recent years has been the polymerase chain reaction or PCR.9  One of the reasons for the adoption of the PCR is the elegant simplicity of the reaction and relative ease of the practical manipulation steps. Frequently this is one of the first techniques used when manipulating DNA and has opened up the analysis of numerous cellular and molecular events in many disease processes (Table 1.2).

Table 1.2

Examples of applications of PCR in various fields of biosciences. Reproduced from ref. 22 with permission from the Royal Society of Chemistrya

Field of study Applications Specific uses
DNA amplification  General molecular biology  Screening gene libraries 
Bridge PCR  Next-generation sequencing  Template cluster preparation 
Production/labelling  Gene probe production  Use with blots and hybridisations 
Reverse transcriptase PCR (RT-PCR)  RNA analysis  Active latent viral infections 
Scenes of crime  Forensic science  Analysis of DNA from blood 
Microbial detection  Infection and disease monitoring  Strain typing and analysis (RAPDs) 
Cycle sequencing  Sequence analysis  Rapid DNA sequencing 
Referencing points in a genome  Genome mapping studies  Sequence tagged sites (STS) 
mRNA analysis  Gene discovery  Expressed sequence tags(ESTs) 
Detection of known mutations  Genetic mutation analysis  Screening for cystic fibrosis 
Digital PCR (dPCR)  Quantification  Viral copy number analysis 
Quantitative PCR (qPCR)  Quantification analysis  5′ nuclease (TaqMan assay) 
Detection of unknown mutations  Genetic mutation analysis  Gel-based PCR methods (DGGE) 
Production of novel proteins  Protein engineering  PCR mutagenesis 
Retrospective studies  Molecular archaeology  Dinosaur DNA analysis 
Sexing or cell mutation sites  Single-cell analysis  Sex determination of unborn 
Studies on frozen sections  In situ analysis  Localisation of DNA and/or RNA 
Field of study Applications Specific uses
DNA amplification  General molecular biology  Screening gene libraries 
Bridge PCR  Next-generation sequencing  Template cluster preparation 
Production/labelling  Gene probe production  Use with blots and hybridisations 
Reverse transcriptase PCR (RT-PCR)  RNA analysis  Active latent viral infections 
Scenes of crime  Forensic science  Analysis of DNA from blood 
Microbial detection  Infection and disease monitoring  Strain typing and analysis (RAPDs) 
Cycle sequencing  Sequence analysis  Rapid DNA sequencing 
Referencing points in a genome  Genome mapping studies  Sequence tagged sites (STS) 
mRNA analysis  Gene discovery  Expressed sequence tags(ESTs) 
Detection of known mutations  Genetic mutation analysis  Screening for cystic fibrosis 
Digital PCR (dPCR)  Quantification  Viral copy number analysis 
Quantitative PCR (qPCR)  Quantification analysis  5′ nuclease (TaqMan assay) 
Detection of unknown mutations  Genetic mutation analysis  Gel-based PCR methods (DGGE) 
Production of novel proteins  Protein engineering  PCR mutagenesis 
Retrospective studies  Molecular archaeology  Dinosaur DNA analysis 
Sexing or cell mutation sites  Single-cell analysis  Sex determination of unborn 
Studies on frozen sections  In situ analysis  Localisation of DNA and/or RNA 
a

Abbreviations: RT, reverse transcriptase; RAPDs, rapid amplification polymorphic DNA; DGGE, denaturing gradient gel electrophoresis.

PCR is used to amplify a precise fragment of DNA (the target) from a complex mixture of starting material, usually termed the template, which may be DNA from microbes, mouth swabs, blood, urine, tissue biopsy.10  It does require the knowledge of some DNA sequence that flanks the fragment of DNA to be amplified (target DNA). From this information two oligonucleotide primers may be chemically synthesised each complementary to a stretch of DNA to the 3′ side of the target DNA, one oligonucleotide for each of the two DNA strands (Figure 1.11) that may be subsequently extended by a DNA polymerase.

Figure 1.11

The location of PCR primers. PCR primers are designed based on sequences adjacent to the region to be amplified, allowing a region of DNA (e.g. a gene) to be amplified from a complex starting material of genomic template DNA. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.11

The location of PCR primers. PCR primers are designed based on sequences adjacent to the region to be amplified, allowing a region of DNA (e.g. a gene) to be amplified from a complex starting material of genomic template DNA. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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One of the reasons for the global adoption of PCR is the elegant simplicity of the reaction and relative ease of the practical manipulation steps. Importantly, the process, which requires a cyclic process of heating and cooling of the individual reagents, is automated. The reagents are added to sample tubes with thin walls for efficient thermal transfer or to 96-well microtitre plates, although there are various formats available. In many cases most of the reagents are pre-mixed in what is termed a ‘master mix’, this minimises pipetting errors and is important since small volumes are usually used. The tubes are then placed in an automated thermal cycler. There are numerous makes and models of different configurations on the market that employ various methods for heating and cooling, the most common uses Peltier effect heating blocks and water or fan cooling. High-end thermal cyclers may also include a robotic sample preparation system and many can be programmed directly on the cycler or through a connected PC.

One problem with early PCR reactions was that the temperature needed to denature the DNA also denatured the DNA polymerase. However, the availability of a thermostable DNA polymerase enzyme isolated from the thermophilic bacterium Thermus aquaticus (found in hot springs) provided the means to automate the reaction. Taq DNA polymerase has a temperature optimum of 72 °C and survives prolonged exposure to temperatures as high as 96 °C and so is still active after each of the denaturation steps. For many applications PCR has replaced the traditional DNA cloning methods as it fulfils the same function, the production of large amounts of DNA from limited starting material; however, this is achieved in a fraction of the time needed to clone a DNA fragment.

PCR consists of three defined steps which are repeated a defined number of times or cycles. In the first cycle the double stranded ‘high molecular weight’ template DNA is denatured by heating the reaction mix to above 90 °C. Within the complex mass of DNA strands, the target region to be specifically amplified is thus made accessible and is single stranded. The temperature is then cooled to between 40 and 60 °C to allow the hybridisation of the two oligonucleotide primers, which are present in excess, to bind to complementary sites that flank the target DNA. This annealing step is unique for each PCR and thus has a defined temperature. The annealed oligonucleotides then act as primers or start points for DNA synthesis since they provide a free 3′ hydroxyl group for DNA polymerase. The DNA synthesis step is termed extension and carried out at 72 °C by a thermostable DNA polymerase, most commonly Taq DNA polymerase (Figure 1.12).

Figure 1.12

The steps involved in one cycle of the polymerase chain reaction. Denaturation of the template strands is followed by annealing of the primers to the template after which extension by Taq polymerase from the 3′ end of the primers takes place. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.12

The steps involved in one cycle of the polymerase chain reaction. Denaturation of the template strands is followed by annealing of the primers to the template after which extension by Taq polymerase from the 3′ end of the primers takes place. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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DNA synthesis proceeds from both of the primers until the new strands have been extended along and beyond the target DNA to be amplified. It is important to note that, since the new strands extend beyond the target DNA they will contain a region near their 3′ ends which is complementary to the other primer (Figure 1.13). Thus, if another round of DNA synthesis is allowed to take place not only the original strands will be used as templates but also the new strands. The products obtained from the new strands will have a precise length, delimited exactly by the two regions complementary to the primers. As the system is taken through successive cycles of denaturation, annealing and extension all the new strands will act as templates and so there will be an exponential increase in the amount of DNA produced. This is against a background of linear amplification of the long original template strands. The net effect is to selectively amplify the target DNA and the primer regions flanking it, leading to the production of millions of identical copies.

Figure 1.13

Terms associated with amplification components and resulting PCR products. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.13

Terms associated with amplification components and resulting PCR products. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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For efficient annealing of the primers, the precise temperature at which the annealing occurs is critical and each PCR system has to be defined and optimised by the user.11  One useful technique for optimisation of annealing temperature is called touchdown PCR where a specialised programmable thermocycler is used to incrementally decrease the temperature until the optimum annealing temperature is reached. Reactions that are not optimised may give rise to other misprimed DNA products in addition to the specific target or may not produce any amplified products at all. Another approach to reducing spurious non-specific amplification during the early stages of the reaction is termed ‘hot start’ PCR. In this method the reaction components are heated to the melting temperature before adding the polymerase. At one time hot start was achieved by introducing a physical wax barrier between the Taq polymerase and the remainder of the reaction components. The wax melted at the denaturation temperature allowing the Taq polymerase access to the reaction mix. More recently, modified polymerase enzyme systems have been developed that inhibit polymerisation at ambient temperature, either by the use of binding ligands, such as antibodies, or by the presence of bound inhibitors that dissociate only after a high-temperature activation step is performed. Taq polymerase has a number of limitations, such as the lack of proof reading resulting in misincorporation errors (some as high as 1 × 10−5 errors per base) and sensitivity to inhibitors, such as those found in blood e.g. haem, immunoglobulin M, or in extraction buffers (proteinase K, phenol etc.). It also leaves a residue at the end of the amplicon which can be useful for dA:dT cloning (see Chapter 3). Engineered versions of the enzyme, which overcome some of these problems, have been produced and the use of other enzymes, such as Pfu (from Pyrococcus furiosus) DNA polymerase, which have proof-reading ability, or combinations of enzymes can be especially useful when amplifying fragments of 5 kb or longer. Further cocktails of DNA polymerases can be used in the reaction to amplify PCR products of 20–30 kb. There have been a number of enhancements for the PCR, typically they try to improve the accessibility of the polymerase to the template, chemicals such as betaine, dimethyl sulphoxide (DMSO), T4 gene 32 protein and single-stranded binding protein (SSBP). One interesting approach is nanoparticle PCR or nanoPCR, where adding gold nanoparticles at nanomolar concentrations can increase the sensitivity of the reaction by as much as a 1000-fold. This may be due to a number of factors, including the enhanced thermal transfer properties of the particles and/or assisting with primer–template binding. Indeed the potential use of nanoparticles is changing the way PCR may be undertaken in the future and with increasing use of carbon nanotubes and quantum dots the specificity and sensitivity may be improved.12 

The specificity of the PCR lies in the design of the two oligonucleotide primers.13  These are required to be complementary to sequences flanking the target DNA but must not be self complementary and form hairpins or inadvertently bind each other to form dimers, since both prevent authentic DNA amplification. They also have to be matched in their GC content, have similar annealing temperatures and be incapable of mispriming and therefore amplifying unwanted genomic sequences. Manual design of primers is time consuming and often hit or miss, although formulae such as the following are still used to derive the annealing temperature or Ta for each primer:

where Tm is the melting temperature of the primer–target duplex and G, C, A and T are the numbers of the respective bases in the primer. In general the Ta is set 3–5 °C lower than the Tm. On occasions, secondary or primer dimer bands may be observed on the electrophoresis gel in addition to the authentic PCR products. In these situations touchdown or hot-start regimes may help. Alternatively, increasing the Ta closer to the Tm can improve the specificity of the reaction. The increasing use of bioinformatics resources, such as primer3, primer-blast at the National Center for Biotechnology Information (NCBI) and primer design assistant, in the design of primers makes the design and the selection of reaction conditions much more straightforward (see Chapter 5). These computer-based resources allow the sequences to be amplified, primer length, product size, GC content etc. to be input and, following analysis, provide a choice of matched primer sequences. Indeed the initial selection and design of primers without the aid of bioinformatics would now be unnecessarily time-consuming. Some of these methods are useful when designing a multiplex PCR where more than one set of primers is included in a PCR. Here multiple amplicons are produced but the cycling conditions and the primer sets require careful design and optimisation. Bioinformatic resources, such as primerplex, are also able to assist with the process of multiplex PCR design. Finally, before ordering or synthesising the primers it is wise to submit proposed sequences to a nucleotide sequence search program, such as Blast, which can be used to interrogate GenBank or other comprehensive public DNA sequence databases to increase confidence that the reaction will be specific for the intended target sequence.

RT-PCR is an extremely useful variation of standard PCR which enables the amplification of specific mRNA transcripts from biological samples without the need for the rigorous extraction procedures associated with mRNA purification which are required for conventional cloning purposes.14  Conveniently, the dNTPs, buffer, Taq polymerase, oligonucleotide primers, reverse transcriptase (RT) and the RNA template are added together to the reaction tube. The reaction is heated to 37 °C, thus allowing the RT to work and enables the production of a cDNA copy of the RNA strands that anneal to one of the primers in the mix. Some thermostable DNA polymerases such as Tth (from Thermus thermophilus) have a reverse transcriptase activity when manganese is added to the reaction buffer. Thus a single step RT-PCR can be undertaken rather than two-step reaction as in the traditional method.

Following ‘first strand synthesis’ a standard PCR is carried out to amplify the cDNA product, resulting in ‘second strand synthesis’ and subsequently a double-stranded DNA (dsDNA) product. The choice of primer for the first strand synthesis depends on the experiment. If amplification of all mRNAs in the cell extract is required then an oligo dT primer that would anneal to all the poly-A tails can be used. If a specific cDNA is sought, then a coding-region-specific primer can be used. Alternatively, random hexamers may be used as described in Section 1.4.6 (Figure 1.14). RT-PCR has many applications, such as the assessment of transcript levels in different cells and tissues when analysed with quantitative PCR (qPCR), is widely used as a diagnostic tool and is especially useful in the fields of microbiology and virology as indicated in Table 1.2. A recent development in this area to try to quantitate rare alleles or viral nucleic acid copies is termed digital PCR (dPCR). Here the template is diluted or partitioned out to millions of single strands distributed in miniature chambers, emulsions and other surfaces. A PCR reaction is then performed such that a positive (1) or negative (0) will result. The determination of how many copies were in the original sample can then be performed. Here technological improvements have allowed the partition and amplification in titrated emulsions of oil, which lowers the cost in comparison with other quantitative methods, such as qPCR. This approach of solid-phase PCR is also used in the template preparation for some of the next-generation sequencing methods.15 

Figure 1.14

Reverse transcriptase PCR (RT-PCR). In RT-PCR mRNA is converted to complementary DNA (cDNA) using the enzyme reverse transcriptase. The cDNA is then used directly in the PCR. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.14

Reverse transcriptase PCR (RT-PCR). In RT-PCR mRNA is converted to complementary DNA (cDNA) using the enzyme reverse transcriptase. The cDNA is then used directly in the PCR. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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The exquisite sensitivity of the PCR system is also one of its main drawbacks since the very large degree of amplification may make the system vulnerable to contamination. Even a trace of foreign DNA or previously amplified patient DNA, even such as that present on laboratory bench surfaces, may be amplified to significant levels and may give misleading results. Hence, cleanliness is paramount when undertaking PCR, and dedicated equipment and, in some cases, dedicated laboratories are used. It is possible that amplified products may also contaminate the PCR although this may be overcome by UV irradiation to damage already amplified products so that they cannot be used as templates. A further solution is to incorporate uracil into the PCR and then treat the products with the enzyme uracil n-glycosylase (UNG) which degrades any PCR amplicons with incorporated uracil, rendering them useless as templates. This is especially important when analysing patient clinical samples. It is of course vital to include the correct negative and positive controls for each PCR to allow subsequent troubleshooting to be undertaken.

One important evolution of the PCR method has been the development of quantitative PCR (qPCR). With this method it is possible to determine initial amounts of starting template DNA and follow the reaction in real time. It also provides an early indication of any problems with the amplification process. Early quantitative PCR methods involved the comparison of a standard or control DNA template amplified with separate primers at the same time as the specific target DNA.16  These types of quantification rely on the reaction being exponential and so any factors affecting this may also affect the result. Other methods have involved the incorporation of a radiolabel through the primers or nucleotides and its subsequent detection following purification of the amplicon.

The introduction of thermal cyclers that incorporate the ability to detect the accumulation of DNA through fluorescent dyes binding to the DNA has rapidly transformed this area.17  One common DNA binding dye is SYBR green, this binds to dsDNA as it is amplified, albeit in a non-specific fashion, and may be detected in a qPCR thermal cycler. In order to quantitate unknown DNA templates a standard dilution is prepared using template DNA of known concentration. As the DNA accumulates during the early exponential phase of the reaction an arbitrary point is taken where each of the diluted DNA samples cross a line. This is termed the crossing threshold or Ct value. From the various Ct values a log graph is prepared from the standards and thus any unknown amplified at the same time can be deduced. Since SYBR green and similar DNA-binding dyes are non-specific, in order to determine if a correctly sized PCR product is present most qPCR cyclers have a built-in melting curve function. This increases the temperature of the amplicons in the cycler until the dsDNA is denatured. Each amplicon has a characteristic melt temperature and thus can be identified on the basis of theoretical and previous data. A refinement of this method, termed precision melt analysis, is able to differentiate between amplicons with one base difference and has value in mutation analysis. Subsequent DNA sequencing can provide a definitive confirmation.

In order to detect specific amplicons during PCR an oligonucleotide probe labelled with a fluorescent reporter and quencher molecule at either end can included in the reaction in the place of SYBR green. This method is termed the 5′ fluorogenic exonuclease detection system or more commonly the TaqMan system. When the oligonucleotide ‘TaqMan’ probe binds to the target sequence the 5′ exonuclease activity of Taq polymerase degrades and releases the reporter from the quencher during extension. A signal is thus generated which increases in direct proportion to the number of starting molecules. Thus the detection system is able to induce and detect fluorescence in real time as the PCR proceeds. Importantly, it provides confirmation that the correct DNA sequence has been amplified (Figure 1.15). Refinements of the PCR process are also in development and promise rapid amplification and reporting. This is mainly due to advances in miniaturising the amplification system and using microfluidics on a small chip. Efficiency, multiplex throughput and low reagent costs are advantages of these methods. Indeed, there are a number of further developments where PCR and other systems including isothermal cycling systems, such as loop-mediated isothermal amplification (LAMP), can be undertaken in various microfluidic platforms. At present, there is a move to provide a set of guidelines or information in the development and reporting of qPCR termed MIQE guidelines or minimum information for publication of quantitative real-time PCR experiments.18  Although not without its drawbacks PCR is a remarkable development that has changed the approach of many researchers to the analysis of nucleic acids and continues to have a profound effect on core genomic and genetic analysis.

Figure 1.15

Representation of the 5′ nuclease assay or TaqMan approach of specific quantitative PCR where R is the reporter and Q is the quencher. Note: Taq polymerase extends from the 3′ end of the primer in the middle panel and cleaves the R from Q in the lower panel, producing a fluorescent signal. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

Figure 1.15

Representation of the 5′ nuclease assay or TaqMan approach of specific quantitative PCR where R is the reporter and Q is the quencher. Note: Taq polymerase extends from the 3′ end of the primer in the middle panel and cleaves the R from Q in the lower panel, producing a fluorescent signal. Reproduced from ref. 22 with permission from the Royal Society of Chemistry.

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Many traditional methods in molecular biology have now been superseded by PCR and the applications of the technique appear to be unlimited. The success of the PCR process has given impetus to the development of other amplification techniques that are based on either thermal cycling or non-thermal cycling (isothermal) methods. Indeed the development of isothermal systems, such as the LAMP DNA amplification system, obviates the need for a thermal cycler and has the advantage of being able to be used in non-laboratory settings.19 

Gross chromosomal changes are often detectable by microscopic examination of the chromosomes within a karyotype. Single or restricted numbers of base substitutions, deletions, rearrangements or insertions are far less easily detectable but may induce similarly profound effects on normal cellular biochemistry. In situ hybridisation makes it possible to determine the chromosomal location of a particular gene fragment or gene mutation. This is undertaken by preparing a labelled DNA or RNA probe and applying this to a tissue or chromosomal preparation fixed to a microscope slide. Any probe that does not hybridise to complementary sequences is washed off and an image of the distribution or location of the bound probe is viewed with a detection system. Using tissue or cells fixed to slides it is also possible to carry out in situ PCR and qPCR. This is a highly sensitive technique where PCR is carried out directly on the tissue slide with the standard PCR reagents. Specially adapted thermal cycling machines are required to hold the slide preparations and allow the PCR to proceed. This allows the localisation and identification of, for example, single copies of intracellular viruses and, in the case of qPCR, the determination of initial concentrations of nucleic acid.

There are several types of mutations that can occur in nucleic acids, either transiently or stably incorporated into the genome. During evolution, mutations may be inherited in one or both copies of a chromosome, resulting in polymorphisms within the population. Mutations may occur at any site within the genome; however, there are several instances whereby mutations occur in limited regions. This is particularly obvious in prokaryotes, where elements of the genome (termed hypervariable regions) undergo extensive mutations to generate large numbers of variants, by virtue of the high rate of replication of the organisms. Similar hypervariable sequences are generated in the normal antibody immune response in eukaryotes. Mutations may have several effects upon the structure and function of the genome. Some mutations may lead to undetectable effects upon normal cellular functions, termed conservative mutations. Examples of these are mutations that occur in intron sequences and therefore play no part in the final structure and function of the protein or its regulation. Alternatively, mutations may result in profound effects upon normal cell function, such as altered transcription rates, or on the sequence of mRNAs necessary for normal cellular processes.

Mutations occurring within exons may alter the amino acid composition of the encoded protein by causing amino acid substitution or by changing the reading frame used during translation. These point mutations were traditionally detected by Southern blotting or, if a convenient restriction site was available, by restriction fragment length polymorphism (RFLP). However, PCR has been used to great effect in mutation and polymorphism detection and numerous techniques have been developed over the last few years.20  These involve both screening and confirmatory techniques. One such is single-strand conformational polymorphism (SSCP), this is a screening method and is able to identify that a mutation is within the amplified region even though the identity of the mutation cannot be determined (see Chapter 2). A further confirmatory PCR-based technique is allele-specific oligonucleotide PCR (ASO-PCR) where two competing primers and one general primer are used in two separate PCRs. One of the primers is directly complementary to the known point mutation whereas the other is a wild-type primer; that is, the primers are identical except for the terminal 3′ end base. Thus, if the DNA contains the point mutation only the primer with the complementary sequence will bind and be incorporated into the amplified DNA, whereas if the DNA is normal the wild-type primer is incorporated. The results of the PCR are analysed by agarose gel electrophoresis. A further modification of ASO-PCR has been developed where the primers are each labelled with a different fluorochrome. Since the primers are labelled differently a positive or negative result is produced directly without the need to examine the PCRs by gel electrophoresis.

Various modifications now allow more than one PCR to be carried out at a time (multiplex PCR), and hence the detection of more than one mutation is possible at the same time. Where the mutation is unknown it is also possible to use a PCR system with a gel-based detection method termed denaturing gradient gel electrophoresis (DGGE). In this technique a sample DNA heteroduplex containing a mutation is amplified by the PCR, which is also used to attach a GC-rich sequence (GC clamp) to one end of the heteroduplex. The mutated heteroduplex is identified by its altered melting properties through a polyacrylamide gel which contains a gradient of denaturant, such as urea. At a certain point in the gradient the heteroduplex will denature relative to a perfectly matched homoduplex and thus may be identified. The GC clamp maintains the integrity of the end of the duplex on passage through the gel. The sensitivity of this and other mutation-detection methods has been substantially increased by the use of PCR, and further techniques used to detect known or unknown mutations are listed in Table 1.3. An extension of this principle is used in a number of detection methods employing denaturing high-performance liquid chromatography (dHPLC). Commonly known as wave technology, the detection of denatured single strands containing mismatches is rapid, allowing a high-throughput analysis of samples to be achieved.

Table 1.3

Methods for detecting mutations in DNA samples. Reproduced from ref. 22 with permission from the Royal Society of Chemistry

Technique Basis of method Main characteristics of detection
Southern blotting  Gel- or blot-based  Labelled probe hybridisation to DNA 
Dot and slot blotting  Blot-based  Labelled probe hybridisation to DNA 
Allele-specific oligo-PCR (ASO-PCR)  PCR-based  Oligonucleotide matching to DNA sample 
Denaturing gradient gel electrophoresis (DGGE)  Gel- or-PCR-based  Melting temperature of DNA strands 
Single-stranded conformation polymorphism (SSCP)  Gel- or PCR-based  Conformation difference of DNA strands 
Multiplex ligation-dependent probe amplification (MLPA)  Gel- or PCR-based  Oligonucleotide matching to DNA sample or ligation 
DNA sequencing  Sanger or next-generation sequencing  Nucleotide sequence analysis of DNA 
DNA microchips  Glass-chip-based  Sample DNA hybridisation to oligo arrays 
Technique Basis of method Main characteristics of detection
Southern blotting  Gel- or blot-based  Labelled probe hybridisation to DNA 
Dot and slot blotting  Blot-based  Labelled probe hybridisation to DNA 
Allele-specific oligo-PCR (ASO-PCR)  PCR-based  Oligonucleotide matching to DNA sample 
Denaturing gradient gel electrophoresis (DGGE)  Gel- or-PCR-based  Melting temperature of DNA strands 
Single-stranded conformation polymorphism (SSCP)  Gel- or PCR-based  Conformation difference of DNA strands 
Multiplex ligation-dependent probe amplification (MLPA)  Gel- or PCR-based  Oligonucleotide matching to DNA sample or ligation 
DNA sequencing  Sanger or next-generation sequencing  Nucleotide sequence analysis of DNA 
DNA microchips  Glass-chip-based  Sample DNA hybridisation to oligo arrays 

Polymorphisms are particularly interesting elements of the human genome and as such may be used as the basis for differentiating between individuals. All humans carry repeats of sequences known as minisatellite DNA, of which the number of repeats varies between unrelated individuals. Hybridisation of probes which anneal to these sequences using Southern blotting provides the means to type and identify those individuals.

Multiplex ligation-dependent probe amplification (MLPA) is a PCR-based multiplex assay that allows a number of target sites, such as deletions, duplications, mutations or single-nucleotide polymorphisms (SNPs) to be amplified with a pair of primer-containing probes. The process involves a denaturation and hybridization stage, a ligation stage and a PCR amplification stage, followed by analysis of the products. The technique involves designing two adjacent hybridization probes that contain the fluorescently labelled forward and reverse primer sequence used the amplification stage. One of the probes is designed with a stuffer sequence that can be varied to suit the target. When binding to the target, the probes are first hybridized to the denatured sample DNA. In the next stage the two oligonucleotide probes are ligated with a ligase, only ligated probes can be used in the amplification stage. Oligonucleotide probes that are not ligated will only contain one primer sequence and, as a consequence, cannot be amplified and generate a signal.

An alternative labelling strategy used in karyotyping and gene localisation is fluorescent in situ hybridisation (FISH).21  This method, sometimes termed chromosome painting, is based on in situ hybridisation but different gene probes are labelled with different fluorochromes, each specific for a particular chromosome. The advantage of this method is that separate gene regions may be identified and comparisons made within the same chromosome preparation. The technique is also likely to be highly useful in genome mapping for ordering DNA probes along a chromosomal segment.

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