Details of the PCR Process

PCR requires a thermostable DNA polymerase, (deoxy)nucleotides of each base (collectively referred to as dNTPs), the target sequence, and a pair of oligonucleotides (referred to as primers) complementary to opposite strands flanking the sequence to be amplified (the target sequence). In the first step, target duplexes are denatured into single strands by heat (Figure 17-2). When the mixture is cooled, primers provided in great excess (usually over a million times the concentration of the initial target) specifically anneal to complementary sequences on the target. Once the primers are annealed, the action of the polymerase synthesizes two new DNA strands by extending each of the two primers at their 3′ end. The primers are designed to recognize sequences of the target that are close enough together that the polymerase extends each strand far enough to include the priming site of the other primer. Usually, the optimal temperature for polymerization (or DNA synthesis) is an intermediate temperature between the annealing temperature (at which primer hybridizes to its target) and the denaturation temperature (at which the newly generated DNA strand dissociates from its template). The second cycle also begins with denaturation, but now twice as many strands (the original genomic DNA and the extension products from the first cycle) are available for primer annealing and subsequent extension. Temperature cycling (typically) employs three temperatures: a high temperature sufficient to denature the target sequence, a low temperature that allows annealing of the primers to the target, and a third temperature optimal for polymerase extension. The instrument that takes samples through the multiple steps of changing temperature is known as a thermocycler.

Repetitive thermocycling results in the exponential accumulation of the short product (consisting of primers and all intervening sequences). If the efficiency of each cycle is optimal, the number of target sequences doubles each cycle (the efficiency is 100% or 1.0). PCR efficiency depends on the primers and the temperature-cycling conditions, along with the presence or absence of polymerase inhibitors. Amplified products accumulate exponentially in the beginning cycles of PCR. At some point, however, the efficiency of amplification falls and eventually the amount of product plateaus (Figure 17-3) as the result of exhaustion of components or competition between primer and product annealing (i.e., the single strands of product are at such high concentrations that they anneal to each other rather than to the primers). The S-curve shape is similar to the logistical model for population growth. In a typical PCR reaction using 0.5 µmol/L of each primer, the maximum DNA concentration achievable is about 100 billion copies/µL.

With the addition of an initial reverse transcriptase step to form cDNA from RNA in the sample, RNA targets can be successfully amplified into DNA copies. Reverse transcription and DNA amplification steps usually are catalyzed by two different polymerases, but some thermostable enzymes (such as the Tth polymerase) have both DNA polymerase and reverse transcription activities, so that both steps can be performed in the same tube with the same enzyme.

After amplification, the products can be detected by various methods. Gel electrophoresis with ethidium bromide staining is a classical qualitative method that may suffice for many applications. When greater accuracy is required, one of the primers can be fluorescently labeled so that after PCR, the fragments can be sized on a DNA sequencing device. Alternatively, some form of hybridization assay can be used to verify or analyze the amplified product. Automated methods are always attractive, and closed-tube methods are particularly advantageous in the clinical laboratory. Adding a fluorescent dye or probe before amplification allows thermocyclers equipped with optical detection to analyze the reaction as it progresses (real-time PCR) or after the reaction is complete (end point melting) without the need to process the sample for a separate analysis step.

PCR Kinetics and Rapid Cycling

It is natural to think about PCR in terms of three events—denaturation of double-stranded target, annealing of primers to their targets, and extension of the DNA strand from the primer—that occur at three temperatures, each requiring a certain amount of time. Indeed, it is common to perform PCR by holding the reaction mixture at three different temperatures (e.g., denaturation at 94 °C, annealing at 55 °C, and extension at 72 °C). Standard thermocycling instruments that use conical tubes focus on accurate temperature control of the heating block at equilibrium, not on the dynamic control of the sample temperature. As a result, sample temperatures are not well defined during transitions, and long cycle times have become standard to ensure that samples reach target temperatures. Reproducibility between instruments and manufacturers is poor, and PCR may require hours to complete 30 cycles of amplification.

The kinetics of PCR suggests that rapid transitions between temperatures with minimal or no pauses (temperature plateaus) provide a better paradigm of PCR amplification (Figure 17-4). Denaturation, annealing, and extension are very rapid reactions as shown by experiments in capillaries.¹²⁵ The use of temperature “spikes” at denaturation and annealing, instead of extended temperature plateaus, allows for rapid cycling with the appropriate instrumentation.¹²⁹ The actual time required for PCR depends on the size of the product, but when it is less than 500 base pairs (bp), 30 cycles are easily completed in 15 minutes. Furthermore, rapid amplification improves specificity. Figure 17-5 shows PCR amplification of the same product amplified at different cycling speeds. With conventional slow cycling, many nonspecific products are generated (cycling profile A). These products disappear as the cycling time is decreased (profiles B, C, and D). In fact, amplification yield and product specificity are optimal when denaturation and annealing times are minimal.

Requirements for denaturation, annealing, and extension in PCR have been reviewed.¹²⁸ Initial denaturation of genomic DNA may be required before PCR cycling, depending on how the DNA sample was prepared. Boiling of the sample or an initial denaturation step of 5 to 10 seconds before PCR cycling may be necessary. During PCR, however, denaturation occurs very rapidly. Even for long PCR products, denaturation is complete in less than 1 second after the denaturation temperature is reached. Anything greater than a denaturation time of “0” serves only to degrade the polymerase. If longer denaturation times are required, either the sample is not reaching temperature or heat-activated polymerases are being used. Product specificity is optimal when annealing times are less than 1 second. Longer annealing times may be required if the primer concentrations are low. The required extension time for each cycle depends on the length of the PCR product. Extension is not instantaneous, although it is much faster than common practice would suggest. Extension rates of Taq polymerase under optimal conditions are 50 to 100 bases per second. Despite its common use, a 5 to 10 minute final extension is not necessary to complete extension. Products can be as small as 40 bp to about 40 kilobases (kb). To amplify products longer than 2 kb, mixtures of polymerases that include some 3′-exonuclease activity to edit mismatched nucleotides are usually used. Instead of separate annealing and extension temperatures, both processes can be carried out at the same temperature, resulting in two-temperature, instead of three-temperature, cycling. Although this simplifies the demand on instrumentation and programming, it limits the choice of primers and requires a longer extension time at suboptimal temperatures.

PCR Optimization and Primer Design

In addition to the temperature-cycling conditions, the specificity of PCR depends on the choice of primers and the Mg²⁺ concentration. The choice of primers often dictates the quality and success of the amplification reaction. To select primers, the sequence of the target must be known. Some guidelines for primer selection are intuitive and helpful:

For exponential PCR, priming sites must be oriented appropriately within a close distance. The desired target can be favored by choosing a small product size and rapid cycling.

Detection Limits of PCR

When PCR is performed under optimal conditions, a single copy of the target can be detected. In practice, however, the statistical probability of distributing at least a single copy from a dilute template solution into the PCR must be considered. The Poisson distribution indicates that if, on average, one target copy is present per tube, 37% of the tubes will have no target, 37% will have one target, and the remainder will have more than one target. About three copies on average are necessary for 95% of the tubes to include at least one copy. Therefore, the limit of detection (95% probability) of any single PCR cannot be lower than three copies per reaction. This limitation of low copy analysis holds true for any amplification technique.

3′-End of PCR Products

Taq DNA polymerase and other polymerases have a terminal transferase activity that may add a single unpaired nucleotide on the 3′-end of PCR product strands. In the presence of all four dNTPs, dATP is preferentially added. This means that some percentage of the double-stranded products generated by PCR will have a protruding A at one or both ends. Although this does not influence most detection protocols, it can complicate some systems with high size resolution. On the other hand, this feature is useful for high-efficiency cloning and ligation of PCR products.¹³⁴

Contamination Control to Avoid False-Positive Results

Because PCR can detect a single molecule of target sequence, a small amount of contamination in a sample can easily produce a false-positive result. The greatest potential for contamination comes from the product of the amplification reaction, referred to as the amplicon (used interchangeably with PCR product). After amplification, each reaction mixture may contain as many as one trillion copies of the amplicon. Thus, minute aerosol droplets contain more than enough target for robust amplification. Amplicon can contaminate reagents, pipettes, and glassware. It is easy to turn a laboratory into a Dr. Seuss fiasco.⁹⁵ Experience has dictated the use of laboratory procedures that minimize contamination by amplicons.⁵² These include physically separated areas for preamplification and postamplification steps, positive-displacement or filter barrier pipettes to minimize aerosol contamination, and the use of prealiquoted reagents. The most effective way of all is to not let the product out of the tube. Methods that perform amplification, detection, and characterization in a closed tube virtually eliminate the risk of product contamination. Even with these precautions, a negative control or blank (all reactants minus target DNA) is one of the most important controls for PCR.

In combination with the precautions listed previously, chemical modifications can be made to the amplicon so it becomes an unsuitable template for further amplification. One of the commonly practiced chemical modifications is to substitute deoxyuridine 5′-triphosphate (dUTP) for deoxythymidine 5′-triphosphate (dTTP) in amplification reactions, which results in incorporation of U in place of T in the amplified product. A bacterial enzyme, UNG, degrades DNA that contains U. Because U is not normally found in DNA, only amplicons will be susceptible to degradation by UNG treatment, and a new target DNA will not be affected. During a brief incubation step before amplification, uracil-containing DNA strands that are carried over from previous amplifications are enzymatically degraded such that they cannot serve as substrates for further amplification. UNG is then inactivated during the first denaturation cycle, so newly formed amplicon can accumulate normally during the reaction. However, UNG may regain activity, even after multiple cycles of amplification, if the temperature of the reaction mixture drops to below 55 °C. Residual UNG activity may also affect the detection limit (and the sensitivity of quantitative PCR assays) if amplified products are held at room temperature before detection. Often, PCR is carried out with dUTP but not with UNG until amplicon contamination is suspected.

An inexpensive method for reducing contamination in PCR consists of exposing the amplicon to ultraviolet (UV) light (such as a transilluminator) for a certain amount of time (usually 15 minutes) immediately after the amplification procedure. UV light creates unamplifiable pyrimidine dimers in the amplicon molecules. However, this procedure may not be compatible with all detection methods.

Inhibition Control to Avoid False-Negative Results

PCR is a resilient process that does not require highly purified nucleic acid. In practice, however, clinical samples may contain impurities that can inhibit polymerase activity. To ensure reliable amplification, some form of nucleic acid purification is often used. The idiosyncratic nature of PCR inhibitors contained in clinical specimens requires demonstration that the sample (or preparation of nucleic acid purified from it) will allow amplification. A control nucleic acid sequence, usually different from the target, can be added to the sample (or to the extract from the sample). Failure to amplify this control indicates that further purification of the sample is required to remove inhibitors of PCR.

Hot Start Techniques

PCR sensitivity and specificity can be compromised by the formation of unintended low molecular weight artifacts. This process is initiated before PCR, when the primers, template, and polymerase are all together at temperatures below the annealing temperature of PCR. Even at low temperatures, if a primer momentarily anneals to another primer or to an undesired target region, Taq DNA polymerase may extend the complex. If the extension product, in turn, is primed and extended, then unintended, double-stranded products can be formed (e.g., primer-dimers) that serve as amplification templates throughout the reaction. Primer-dimers can be distinguished from the intended target by their molecular weight or melting temperature, but they also influence the efficiency of the intended amplification and decrease the sensitivity of the assay.

The formation of primer-dimers can be minimized in several ways. All limit the activity of polymerase until the temperature is increased (thus the strategy is often collectively called hot start). One method of hot start involves the use of antibody (or an aptamer) to bind and inactivate the polymerase at room temperature. The binding agent is released upon heating, allowing polymerase activation. Another method uses wax or paraffin to create a physical barrier between the essential components in the reaction. This barrier may be created by putting some of the reaction components into the bottom of the tube and overlaying them with molten wax. Cooling solidifies the wax, and the missing components (usually the polymerase, or magnesium, which is essential for polymerase activity) can be placed on top. The wax melts when the temperature reaches 60 to 80 °C, and all components are mixed together by convection while the molten wax floats on top and prevents evaporation of the sample. Various commercial wax beads encasing one or more critical components are available. Finally the polymerase, primers, or dNTPs can be chemically blocked so that extension cannot occur until activation by heat, usually requiring an extended initial denaturation period of 10 to 20 minutes.

Asymmetric PCR and Allele-Specific PCR

Conventional PCR uses primers that are present in equal amounts, thereby ensuring that most of the products are double-stranded amplicons. Asymmetric PCR uses different concentrations of the two primers to generate more of one strand than of the other. For instance, the use of primer A at 0.5 µmol/L and primer B at 0.005 µmol/L produces mostly single-stranded DNA extended off the more abundant primer. This is useful for sequencing or making single-stranded probes. Yield of the product, however, may be low. With less extreme ratios (e.g., primer A at 0.5 µmol/L and primer B at 0.2 µmol/L), the yield is mostly preserved, with one strand produced in enough excess to make it more available for probe hybridization.

Another variant method called allele-specific PCR enables preferential amplification of one genetic allele over another. The 3′-end of one primer is placed at the polymorphic site and is extended readily only if it is completely complementary to the target. This strategy is used for distinguishing a gene from its pseudogenes and for genotyping of single-nucleotide polymorphisms (SNPs). Allele-specific PCR can also be used to determine haplotypes.⁶⁶

Single-Molecule PCR or Digital PCR

Conventional PCR averages the amplification results of the many individual template molecules in a reaction. Because of this averaging, certain questions are difficult to answer. For example, quantification of copy number variants and minor allele fractions in the background of wild-type DNA, although possible, is difficult by bulk PCR. It is also difficult to determine whether multiple variants are on the same or different chromosomes. Single-molecule PCR (sometimes also called digital PCR) is a technique that uses a dilute solution of template that is distributed across many reaction compartments such that each of those compartments will have no template or a single molecule of template. Instead of conventional tubes, these compartments may be minute aqueous droplets in a water-in-oil emulsion (emulsion PCR),¹⁰² PCR colonies (polonies) on a thin film of acrylamide gel,⁶⁸ or clusters on the surface of a planar flowcell generated by bridge amplification,⁴ or they may be found on beads with clonally amplified template attached to their surface.^62,96 When amplification is observed in one of the massively parallel reactions, this means that clonal amplification occurred from a single template molecule. By analyzing each of these clonal reactions, it is possible (1) to determine whether multiple sequence variants are carried on one chromosome or split among two chromosomes, (2) to detect hemizygous gene or exon level deletions, and (3) to count minority events such as a mutation in a gene that is present in a very small fraction of cells in the specimen. Single-molecule PCR is used as the first step for many of the high-throughput sequencing methods reviewed later in this chapter.

Transcription-Based Amplification Methods

Transcription-based amplification methods are modeled after the replication of retroviruses. These methods are known by various names, including transcription-mediated amplification (TMA),30,43 nucleic acid sequence–based amplification (NASBA),¹⁵ and self-sustained sequence replication (3SR) assays.³² They amplify their target without temperature cycling (isothermally) and use the collective activities of reverse transcriptase, RNase H, and RNA polymerase. The most widely used is TMA, illustrated in Figure 17-6. Two primers, a reverse transcriptase, and an RNA polymerase are used. The primer complementary to the RNA target has a 5′-tail that includes a promoter sequence for RNA polymerase. This primer anneals to the target RNA and is extended by the reverse transcriptase, creating an RNA/DNA duplex. The RNA strand is degraded by the RNAse H activity of the reverse transcriptase, allowing the second primer to anneal. The reverse transcriptase then extends the second primer to create double-stranded DNA that includes the promoter. RNA polymerase recognizes the promoter and initiates transcription, producing 100 to 1000 copies of RNA. Each strand of RNA then binds and extends the second primer, forming a RNA/DNA hybrid; the RNA in the hybrid is degraded, the promoter primer binds and extends to produce double-stranded DNA that can be transcribed, and the cycle repeats. As in PCR, all reagents are included and amplification is exponential with completion in less than an hour. Unlike PCR, these methods do not require temperature cycling (except for an initial heat denaturation if a DNA template is used). They are particularly advantageous when the target is RNA (e.g., human immunodeficiency virus [HIV] and hepatitis C virus [HCV] in blood bank nucleic acid testing).

Strand Displacement Amplification

Another isothermal amplification technique is strand displacement amplification (SDA).¹¹⁴ After heat denaturation of DNA in the presence of four primers—dCTP, dGTP, dUTP, and a modified deoxynucleotide (dATPαS)—two enzymes are added: an exonuclease-deficient polymerase and a restriction enzyme. The two flanking primers that enter into exponential amplification have a restriction site added to their 5′-end and get nicked by the restriction enzyme, allowing displacement of strands that in turn can be primed, extended, and nicked. Deoxy-ATPαS is used so that restriction sites include a hemiphosphorothioate linkage to allow single-strand nicking, instead of cutting through the double strands.

Loop-Mediated Amplification Methods

The target amplification methods presented so far produce a defined length of RNA or DNA according to primer placement. In contrast, loop-mediated methods produce a wide range of different length concatemers of the target sequence. Primer design includes tails at the 5′-end of some of the primers that are complementary to the target sequence. After replication of complementary strands, these ends become free 3′-ends capable of extension. By forming a loop, these 3′-ends prime and extend the hairpin (self-priming) initiating concatemer formation. After initial DNA denaturation, the process can be isothermal if strand displacement primers are included. In the basic version, two strand displacement primers and two loop-forming primers recognize six segments in the target.⁷⁵ In another version, allele-specific amplification with five primers and one competitive probe recognizes seven segments in the target.¹ In both versions, a variety of product concatemers are formed, and the reactions can be completed in less than 1 hour.

Whole Genome and Whole Transcriptome Amplification

Instead of specific amplification of one target to improve sensitivity, methods that amplify all genomic DNA or mRNAs are useful when the target is in short supply. For example, multiple-displacement amplification uses exonuclease-resistant random hexamers and a highly processive polymerase to amplify DNA nonspecifically.¹⁹ Initial DNA denaturation is not necessary, and the reaction proceeds isothermally. Similarly, messenger RNA can be generically amplified with a poly(T) primer modified with an RNA polymerase promoter.⁸⁰ After reverse transcription, second-strand DNA synthesis, and transcription, antisense RNA is produced. Both whole genome and antisense RNA amplification are also useful as nucleic acid purification methods before amplification or detection.

Branched-Chain DNA: Signal Amplification

Instead of increasing the concentration of target, signal amplification techniques use nucleic acids to magnify the detection signal. The branched-chain DNA (bDNA) method is one of these techniques in common use. The bDNA approach hybridizes the target nucleic acid to multiple capture probes affixed to a microtiter well.¹⁰⁸ This is followed by hybridization to a series of “extender,” “preamplifier,” and amplifier probes. The final, highly branched amplifier probe includes multiple copies of signal-generating enzymes that act on a chemiluminescent substrate to produce light. Nucleotide analogs isoC and isoG (isomers of C and G that are complementary to each other but not to other nucleotides) are often used to increase the specificity of the signaling cascade.

Serial Invasive Amplification: Signal Amplification

When two probes overlap on one target, an “invasive” cleavage reaction can be catalyzed by certain structure-specific nucleases. The cleaved fragment, in turn, can cause invasive cleavage of a secondary probe in the shape of a hairpin. The hairpin probe can be designed as a fluorogenic indicator by using a reporter/quencher pair of dyes that are separated by cleavage. This serial sequence of events (primary invasion and cleavage, followed by secondary invasion and cleavage of an indicator probe) is known as the serial invasive signal amplification reaction.³⁵ After DNA denaturation, cooling, and the addition of enzymes, the reaction is run at a temperature at which both the primary and secondary reactions recycle.

Rolling Circle Amplification: Probe Amplification

If a primer is annealed to a closed circle of DNA in the presence of a processive, displacing polymerase, the complement of the circle will be synthesized over and over again with displacement of the tandem repeats.⁶⁰ If two primers are used in opposite orientation, progressively more complex branches will be formed in an exponential reaction. The rolling circle can be formed by ligation of the two ends of a linear probe on template DNA. Ligation may happen directly, after polymerization through a gap, or after annealing of an additional, allele-specific oligonucleotide.

UV Absorbance

Nucleic acid molecules absorb ultraviolet light maximally at 260 nm owing almost entirely to the constituent bases. This property can be used to measure the nucleic acid content of a solution. DNA double helices have lower UV absorbance than an equivalent number of nucleotide monomers, and when DNA is denatured into single strands (ssDNA) (e.g., by heat or pH), UV absorbance increases.⁶ If a double-stranded DNA preparation is pure, a 50 mg/L solution has an absorbance of 1.0 at 260 nm. RNA has a greater absorbance, so only about 30 mg/L gives an absorbance of 1.0. More precise estimates for oligonucleotides are based on dinucleotide contributions.⁸ It is common to assess the purity of a nucleic acid preparation by its ratio of absorbances at 260 nm and 280 nm (260 : 280 ratio). In contrast to nucleic acids, proteins absorb maximally at 280 nm. A pure preparation of nucleic acid should have a 260 : 280 ratio between 1.7 and 2.0 that depends on base content. Lower values suggest significant protein contamination.

Fluorescent Staining of Nucleic Acids

Although absorbance measurements are simple and precise, they are not sensitive. Fluorescent stains that bind to nucleic acid are 1000 to 10,000 times more sensitive than absorbance measurements. The best known example of a nucleic acid dye is ethidium bromide, a positively charged, intercalating dye for double-stranded DNA and to a lesser extent, single-stranded DNA and RNA. Cyanine dyes are also popular stains for nucleic acids; they do not fluoresce unless they are bound to nucleic acids, thus providing very low background. With the appropriate optics, single molecules of DNA can be visualized with cyanine-based nucleic acid stains.⁷⁹ Nucleic acid dyes can detect DNA and RNA in gels or in solution (such as in real-time PCR).

Radioactivity

The first probes used in nucleic acid detection were radioactively labeled. Radioactive labels are still favored in some research settings because of the sensitivity attained with probes of high specific activity. The most frequently used isotopic labels are ³²P and ³³P, which are incorporated into the probe by enzymatic reactions. Nick translation is a classical method of labeling double-stranded DNA fragments. Nicks are introduced randomly into each strand of DNA by DNase I, and the resulting 3′-OH groups form priming sites for DNA polymerase. Labeled nucleotides are incorporated as the polymerase extends the strand by digesting and removing the unlabeled strand. Another method of labeling double-stranded DNA is random priming, in which the nucleic acid is denatured and is allowed to anneal with short oligonucleotides of random sequence. The 3′-end of an annealed primer forms an initiation site for the DNA polymerase, which incorporates labeled nucleotides using single-stranded regions of the DNA as the template. Labeled nucleic acid can also be synthesized by PCR, or if a labeled RNA is desired, it can be prepared with the use of a double-stranded DNA fragment containing a promoter sequence for RNA polymerase and incorporating radiolabeled ribonucleotides by transcription.

Additional enzymatic reactions are useful for labeling oligonucleotides. T4 polynucleotide kinase may be used to label the 5′-end of an oligonucleotide. Alternatively, terminal deoxynucleotidyl transferase (TdT) can add labeled nucleotides onto the 3′-end in a tailing reaction. No template is required, and the number and type of nucleotides can be controlled by the reaction conditions. A somewhat longer probe than the original oligonucleotide is obtained with additional labeled bases at the 3′-end. The sensitivity that is achieved with radioactive nucleic acid probes is determined by the radioactivity of the label, its extent of incorporation, and the detection time. Radioactively labeled probes have a short shelf life limited by isotopic decay and radiolysis of the nucleic acid. This inherent instability, along with concerns of radioisotope safety and disposal, restricts the use of radioactive probes in the clinical laboratory.

Indirect Probe Detection

The first practical example of nonradioactive probes used a biotin-labeled analog of dUTP.⁵³ Despite the altered steric configuration, this nucleotide is incorporated by both DNA polymerase and terminal transferase. Other functional groups, such as digoxigenin, may also be used as affinity labels through chemical linkage to a dUTP and incorporation into polynucleotides. Alternatively, oligonucleotide probes can be labeled during synthesis with biotin or amino linkers for subsequent attachment to indicator molecules. Labels at the 5′- or 3′-end of the molecule are usually preferred because central modifications may interfere with hybridization.

Biotin and other affinity labels do not generate detectable signals on their own. However, they can initiate signal amplification mediated by high-affinity binding with antibodies, or in the case of biotin, with avidin or streptavidin. These binding molecules can be linked to enzymes—such as alkaline phosphatase, peroxidase, or luciferase—connecting a single target to a single enzyme. Enzyme activity is monitored by detecting catalytic turnover of enzyme substrates that result in colorimetric, fluorescent, or chemiluminescent signals.

Affinity labels can be used to capture and localize targets to an area of a solid support. For example, biotinylated probes can be affixed to a streptavidin-coated surface. After incubation with the target nucleic acid, a second probe is added, which may be directly labeled with fluorescence or conjugated through another affinity label to an enzyme. Any background or nonspecific localization of reagents results in amplification of an undesired signal along with the desired signal, and these methods usually require multiple separation and washing steps to decrease the background.

Electronic detection of nucleic acids is attractive for its simplicity. Hybridization events can be detected by redox indicators that recognize the DNA duplex, or by other hybridization-induced changes in electrochemical parameters, such as conductivity or capacitance.78,107 However, it is difficult to make electronic methods sensitive and specific enough that nucleic acid amplification is not required.

Fluorescent Labels

Advances in oligonucleotide synthesis and fluorescence detection have made fluorescently labeled probes the preferred reporter for nucleic acid analysis. Many fluorescent labels are now available, allowing color multiplexing for applications such as DNA sequencing, fragment length analysis, DNA arrays, and real-time PCR as reviewed later in this chapter. Techniques such as fluorescence polarization, fluorescence resonance energy transfer (FRET), and fluorescence quenching can provide additional detection specificity. Fluorescence polarization can be used to distinguish free from bound label, if the molecular rotation of the probe changes upon binding.⁵¹ Molecular rotation primarily depends on the size of the molecule, so binding of a small probe onto a large target results in a polarization increase that can be measured. FRET techniques depend on the distance between two spectrally distinct fluorescent labels. Two labeled probes may be brought closer together by adjacent hybridization. Alternatively, two labels on the same probe may end up farther apart by hydrolysis or hybridization. Fluorescence quenching or augmentation does not always require FRET. For example, fluorescence may change merely by hybridization of a fluorescent oligonucleotide to its target. The effect depends on the dye and on inherent quenching from G residues in the target and/or probe.^16,72 Alternatively, quenching moieties can be purposely incorporated into probes.^55,106

Restriction Fragment Length Polymorphism

DNA extracted from a cell is extremely long and is usually cut into shorter fragments before electrophoresis to enhance mobility. Restriction endonucleases cut double-stranded DNA into fragments of reproducible size; the same enzyme produces the same fragments in different specimens if they contain the same DNA sequence. If an alteration in the DNA abolishes or creates a cleavage site recognized by the enzyme (or changes the spacing between two cleavage sites), then different sized fragments will be produced. These changes in fragment lengths (or polymorphisms) that result from restriction digestion are called restriction fragment length polymorphisms (RFLPs). However, restriction digestion produces thousands of fragments. To be useful, specific fragments need to be visualized.

Southern and Northern Blotting

Southern blot analysis (Figure 17-9) separates restriction fragments by agarose gel electrophoresis and transfers (blots) them to a membrane for easier manipulation in the following steps and for selective visualization by labeled probes. It reveals polymorphisms in the DNA sequence based on the RFLP profile. After electrophoresis, acid treatment fragments the DNA to make it easier to blot, and base treatment denatures the DNA to single strands for better binding to the membrane and probe hybridization. After neutralization, the DNA is transferred to a nitrocellulose or nylon membrane and immobilized. The membrane is then incubated with one or more probes that hybridize to the fragments of interest. The hybridized probes then are visualized by autoradiography or chemiluminescence. Southern blotting detects RFLPs, including large structural alterations such as deletions, duplications, insertions, and rearrangements. The procedure was named after its inventor, E. M. Southern,¹⁰⁰ and was the first discrimination method with adequate sensitivity and specificity for DNA analysis of single-copy genes in complex genomes. However, it requires large amounts of DNA (10 to 50 µg/lane) and is very labor intensive and time consuming. Today, it has largely been replaced by PCR.

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