Signal Amplification Techniques

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Branch DNA

Branch DNA (bDNA) analysis is a quantitative technique that is used primarily to quantify levels of certain viruses, such as HIV-1 and HCV, in biological samples. The principle of a bDNA assay is shown in Fig. 11. In this example, the target is an RNA virus molecule. The target RNA is liberated from the virus, and capture extender and label extender molecules are allowed to hybridize to the target. Capture extenders and label extenders are single-strand DNA oligonucleotides that have sequence complementarity to regions of the target RNA. The capture extenders also have a region that can hybridize to immobilized DNA oligonucleotides (capture probes) on the wall of a microtiter plate well. If the target RNA is present in

Amplifiers w"' Labeted Prot

Amplifiers w"' Labeted Prot

Label Extender

Capture Extender

Capture Probe

Fig. 11. Branch DNA (bDNA). See text for details.

Preamplifier

Label Extender

Capture Extender

Capture Probe

Fig. 11. Branch DNA (bDNA). See text for details.

the sample being tested, it will bind to the microtiter plate via the capture probes and the capture extenders. The label extenders can also hybridize to another set of oligonucleotides called preamplifier molecules. These, in turn, hybridize to amplifier molecules. The result of these steps is that multiple amplifier molecules are bound to the wall of the microtiter plate if the target RNA is present in the sample being analyzed.

Detection of the amplifier molecules is by the addition of oligonucleotide-linked alkaline phosphatase molecules, which bind to the amplifier molecules. Addition of a chemiluminescent substrate for alkaline phosphatase generates a chemiluminescent signal that is measured with a photodetector. The intensity of the emitted light is proportional to the number of target RNA molecules in the sample.

bDNA is a proprietary technology of Bayer. Commercially available assays include quantitative tests for HIV-1, HCV, and HBV.

Hybrid Capture

The Hybrid Capture system utilizes a monoclonal antibody that selectively recognizes RNA/DNA duplexes. It has been developed to recognize targets such as human papillomavirus (HPV) DNA. The principle of the Hybrid Capture system is shown in Fig. 12. DNA isolated from the specimen (e.g., a cervical swab) is extracted, dena-

Fig. 12. Hybrid capture assay. See text for details.

tured, and then allowed to hybridize to a synthetic RNA probe that is complementary to the target DNA. Hybridization of the probe to the target leads to the formation of a RNA/DNA duplex that is captured by immobilized antibody to the wall of a microtiter plate. This antibody recognizes RNA/DNA duplexes, but not RNA or DNA single-stranded molecules or homoduplexes. A detection antibody, which also recognizes RNA/DNA duplexes and is conjugated to alkaline phosphatase, then binds to the immobilized duplex. Addition of a chemiluminescent substrate results in emission of light in proportion to the number of copies of the target present in the sample. Hybrid Capture is a proprietary technology of Digene (Gaithersburg, MD).

Reverse Transcription

During gene transcription, the usual direction of flow of genetic information is from DNA to RNA. Several enzymes are known that can synthesize DNA using RNA as a template. Such enzymes are called reverse transcriptases, and the process they catalyze is called reverse transcription (RT) because the direction of information flow is the reverse of normal. Because many laboratory techniques, notably PCR and other cloning schemes, are designed to work with DNA as a template, it is a common prerequisite for such techniques to convert a RNA sequence to its cDNA sequence using RT. Examples of applications in which RT is performed include detection or quantification of gene expression, or of RNA viruses such as HIV-1 or HCV.

For gene expression studies, total cellular RNA can be used for RT, or the RNA can be enriched for messenger RNA (mRNA). The latter represents only a small fraction—about 2-5%—of the total cellular RNA but is composed of molecules that are often of the greatest biological interest. mRNA can be purified by passing total cellular RNA over a column containing bound oligo-dT (i.e., a DNA molecule composed of approx 18 dT residues). The poly-A tail present in most mRNAs binds specifically to the oligo-dT, while the other RNA molecules pass through the column. The enriched mRNA can then be eluted from the column.

In order to perform an RT reaction, a reverse transcriptase is needed. Enzymes in clinical laboratory use include Moloney murine leukemia virus (MMLV) and AMV. Other reagents required for a RT reaction are dNTPs, which serve as the building blocks for the complementary DNA molecule, a suitable buffer, the RNA template, and primers to serve as an anchor for new DNA synthesis.

RT Primers

Three types of reverse transcription primer are commonly used:

1. Oligo-dT is used to prime synthesis of cDNA from mRNA selectively. Most mRNA molecules have a 3' tail consisting of poly-A; oligo-dT can hybridize to this. This approach is used to prime synthesis of expressed genes.

2. Random hexamers. These are random sequences composed of six nucleotides that can be used to prime DNA synthesis at many positions in RNA. This results in synthesis of a large number of cDNA molecules. Each mRNA may be represented by multiple different cDNA molecules.

3. Target-specific primers. These are designed to hybridize selectively to an RNA molecule of interest. Under conditions of suitable stringency, only a particular RNA molecule will be reverse transcribed.

MUTATION DETECTION

Different mutation detection strategies are used to detect specific mutations or to find unknown mutations. The mutation that causes sickle cell anemia is the same in every patient with this disease; therefore a method to test for this specific mutation would be used to identify patients carrying the mutation. On the other hand, hundreds of different mutations in BRCA1 and BRCA2 have been associated with hereditary breast cancer. Detection of these in patient samples requires a different laboratory approach. Some approaches are suitable for both kinds of mutation detection. Methods in common use are described here, but it should be noted that there are many reported variations (16).

Methods to Identify Known Mutations Methods Based on PCR

PCR is widely used in mutation detection because of its rapidity and simplicity. The region of the target gene containing the suspected mutation is amplified. Mutation detection can be accomplished in several ways including RFLP analysis, allele-specific amplification, and detection of mutations by allele-specific oligonucleotides.

PCR Restriction Fragment Length Polymorphism Analysis

If a mutation alters a restriction enzyme site, then that mutation can be demonstrated by incubating the PCR product with the restriction enzyme and analyzing the sizes of the resulting fragments. This size determination can be accomplished by several methods. The most commonly used method in clinical laboratories is electrophoresis in a suitable gel, typically agarose or polyacry-lamide, followed by staining of the gel with ethidium bromide and visualizing the fragments. DNA fragments of known size are run in parallel for size comparison. An example of this approach is shown in Fig. 13. Separations and sizing of fragments by capillary electrophoresis is also in reasonably common use. Other techniques in use include determination of fragment size by mass spectrometry.

PCR with Mutagenic Primers

The PCR-restriction enzyme digestion approach is quite useful if a mutation creates or destroys a naturally occurring restriction enzyme site. It can also be used if a mutation does not alter such a site. By using mutagenic primers, it is often possible to create a restriction enzyme site in the PCR product depending on whether the template DNA contains the mutation of interest. The principle of this approach is shown in Fig. 14. One of the primers used for PCR contains one or more nucleotide substitutions toward its 3' end, the end of the primer that is extended by the DNA polymerase during PCR. These substitutions are designed in such a way that the sequence of the PCR product contains (or doesn't contain) a restriction site depending on whether or not the patient sample contains

Fig. 13. PCR RFLP detection of Cys282Tyr in HFE. (A) Schematic of the 390-bp amplicon of HFE encompassing exon 4 and nucleotide 845 (cDNA sequence). Primers A and B are located in flanking intron sequences. The G845A mutation generates a new Rsal recognition site, as shown in the mutant allele. (B) Photograph of ethidium bromide-stained 12.5% polyacrylamide gel demonstrating PCR RFLP analysis for Cys282Tyr in HFE. W denotes water (reagent) controls. M denotes molecular weight markers. Lanes 1, 2, and 3 show undigested 390-bp amplicons from wild-type (CTRL), heterozygous mutant (HET), and homozygous mutant (HOM) individuals. Lanes 1*, 2*, and 3* show Rsal digestion products from the 390-bp amplicons shown in lanes 1, 2, and 3, respectively. Rsal digestion of a wild-type allele yields fragments of 248 and 142 bp. Rsal digestion of a mutant allele yields fragments of 248, 113, and 29 bp. (The 29-bp fragment is not seen.) (Reproduced with permission from ref. 1).

Fig. 13. PCR RFLP detection of Cys282Tyr in HFE. (A) Schematic of the 390-bp amplicon of HFE encompassing exon 4 and nucleotide 845 (cDNA sequence). Primers A and B are located in flanking intron sequences. The G845A mutation generates a new Rsal recognition site, as shown in the mutant allele. (B) Photograph of ethidium bromide-stained 12.5% polyacrylamide gel demonstrating PCR RFLP analysis for Cys282Tyr in HFE. W denotes water (reagent) controls. M denotes molecular weight markers. Lanes 1, 2, and 3 show undigested 390-bp amplicons from wild-type (CTRL), heterozygous mutant (HET), and homozygous mutant (HOM) individuals. Lanes 1*, 2*, and 3* show Rsal digestion products from the 390-bp amplicons shown in lanes 1, 2, and 3, respectively. Rsal digestion of a wild-type allele yields fragments of 248 and 142 bp. Rsal digestion of a mutant allele yields fragments of 248, 113, and 29 bp. (The 29-bp fragment is not seen.) (Reproduced with permission from ref. 1).

Fig. 14. Mutagenic primer usage to create a restriction site. (A, B The sequence of the prothrombin (factor II) gene is shown around the site of the G^A transition that is associated with increased thrombotic risk (indicated by an asterisk). This mutation does not alter a naturally occurring restriction enzyme recognition sequence. The sequence of a mutagenic primer for PCR is shown below the pro-thrombin gene sequence. This primer contains a mispaired base, indicated by a box. This nucleotide alteration becomes incorporated into PCR products with each cycle of PCR. At completion of PCR, the products amplified by the mutagenic primer contain a HindIII site (AAGCTT) if the 20210A mutation was present, but not if the original template did not contain this mutation. The second member of the pair of PCR primers is not shown in this illustration.

Fig. 14. Mutagenic primer usage to create a restriction site. (A, B The sequence of the prothrombin (factor II) gene is shown around the site of the G^A transition that is associated with increased thrombotic risk (indicated by an asterisk). This mutation does not alter a naturally occurring restriction enzyme recognition sequence. The sequence of a mutagenic primer for PCR is shown below the pro-thrombin gene sequence. This primer contains a mispaired base, indicated by a box. This nucleotide alteration becomes incorporated into PCR products with each cycle of PCR. At completion of PCR, the products amplified by the mutagenic primer contain a HindIII site (AAGCTT) if the 20210A mutation was present, but not if the original template did not contain this mutation. The second member of the pair of PCR primers is not shown in this illustration.

the mutation of interest. The mutagenic primer provides part of the restriction site; the patient's DNA template provides the remaining essential sequence to form the complete restriction site.

This approach has some limitations. Altering the sequence of a primer may alter its specificity for binding to the template DNA and therefore its ability to prime the target of interest efficiently or selectively. Following restriction, the size of the PCR product is reduced by approximately the length of the mutagenic primer, which is usually only about 18 or so nucleotides in length. Such a length change may be difficult to detect if it represents a minor alteration in the overall length of the PCR product. Finally, for cer-

5 1 -CAGCTGTTCGTGTTCTATGATC Pairing

5 ' -CAGCTGTTCGTGTTCTATGATC Pairing

...GTCGACAAGCACAAGATACTAC...

Fig. 15. Allele-specific amplification. In the upper part of the diagram the 3' end of a primer forms a complementary base pair with a target DNA sequence. This primer can be extended during PCR. In the lower part of the diagram the 3' end of a primer has a base pair mismatch with the target DNA sequence. This primer cannot be extended. Successful PCR amplification with the primer can therefore be used to detect the presence of one or other sequence in a DNA sample, in this case, the sequence in the upper part of the illustration. The second member of the pair of PCR primers is not shown in this illustration.

tain mutations, it may not be possible to design a mutagenic primer to create a restriction site.

Allele-Specific PCR Amplification

Allele-specific amplification [also called the amplification refractory mutation system (ARMS)] is based on the inability of many DNA polymerases to extend a primer if the base at the 3' end of the primer does not form a perfect match with the corresponding base in the DNA template (Fig. 15). Extension can take place only if a perfect match exists. If one primer is specific for a given mutation at its 3' end, geometric amplification during PCR can occur only if the target DNA contains this mutation. In a typical allele-specific analysis, two PCR reactions are performed in parallel. One uses a primer that is specific for the mutation, and the other uses a primer specific for the corresponding wild-type allele. Both reactions also use a common second primer. Successful amplification of genomic DNA in only one sample indicates that the DNA is homozygous for that allele. Amplification in both reactions indicates heterozygosity for both wild-type and mutant alleles. Because absence of amplification of the target is a possible outcome, it is usual to coamplify a second, unrelated target using a second pair of primers in the same reaction. Amplification of this second target provides assurance that both amplifiable DNA and the PCR reagents were indeed present in the reaction and therefore that absence of amplification of the primary target is not caused by technical artifact.

Fig. 16. Reverse allele-specific oligonucleotide (ASO) principle. Two oligonucleotides are immobilized on a solid phase. Under the right stringency conditions, the PCR product, shown on the right, can hybridize to the probe with which it has perfect sequence complementarity. Detection of hybridization can be achieved using various techniques such as labeling the PCR product with a radioactive isotope and performing autoradiography.

Fig. 16. Reverse allele-specific oligonucleotide (ASO) principle. Two oligonucleotides are immobilized on a solid phase. Under the right stringency conditions, the PCR product, shown on the right, can hybridize to the probe with which it has perfect sequence complementarity. Detection of hybridization can be achieved using various techniques such as labeling the PCR product with a radioactive isotope and performing autoradiography.

Allele-Specific Oligonucleotide Probes

An allele-specific oligonucleotide (ASO) probe is used to detect the presence of mutations by specific hybridization of the probe to amplified DNA. In this technique, the gene of interest is amplified by PCR, denatured, and then applied to a membrane. Genomic DNA can also be used, but the large amounts of genomic DNA required make this somewhat impractical. The ASO probe, which is single-stranded DNA, generally 18-30 nucleotides in length, is allowed to hybridize to the immobilized DNA under stringency conditions such that the probe can hybridize only if there is perfect sequence complementarity with the target DNA. Under the correct stringency conditions, a mismatch of even a single nucleotide within an oligonucleotide probe of approx 20 nucleotides can inhibit hybridization by altering the optimum annealing temperature of the oligonucleotide. To detect hybridization, the probe can be labeled and detected with a variety of systems, e.g., a radioactive label and autoradiography. For each allele to be detected (i.e., wildtype and mutant), a specific probe is used.

A more commonly used variation of the ASO technique is to immobilize the probe on a membrane or other solid support and allow the amplified, denatured DNA to hybridize to the probe (Fig. 16). This approach is known as reverse ASO. It has the advantage that multiple probes, designed to detect different mutations in the same amplified DNA fragment, can be applied to the solid support. This allows for multiple mutations to be detected simultaneously. In this technique, the amplified DNA fragments are labeled so that hybridization can be detected. This approach can be used in microarrays containing hundreds or thousands of immobilized oligonucleotides to detect many mutations.

Oligonucleotide Ligation Assay

An oligonucleotide ligation assay (OLA) utilizes two oligonu-cleotides that anneal to the target DNA in a head-to-tail fashion, as in ligase chain reaction (Fig. 8). The target DNA of interest is first amplified by PCR. If the 3' end of the first oligonucleotide is perfectly hybridized to the target DNA, then it can be ligated by a DNA ligase to the 5' end of the adjacent oligonucleotide. If the target DNA contains a mutation that prevents the 3' end of the first primer from hybridizing at this position, then ligation does not occur. Detection of ligation can be made by several methods, such as observing the change in molecular weight of the ligated oligonu-cleotides by polyacrylamide gel or capillary gel electrophoresis.

Real-Time PCR

Some thermal cyclers allow for continuous monitoring of the production of PCR products as the reaction proceeds during each cycle. This is known as real-time PCR monitoring, and it can be used to perform quantitative PCR or for qualitative identification of genetic variants. The thermal cycler contains the necessary light source, optics, filters, and detectors to perform fluorescence measurements. The chambers in which each PCR is performed have an optically clear segment to enable fluorescence readings to be made.

For quantitative measurements in real-time PCR, two general approaches are available. Nonspecific quantitative monitoring of a PCR product is performed by observing the fluorescence of a double-stranded DNA-binding dye such as SYBR® Green I. This dye fluoresces in proportion to the amount of DNA in a reaction vessel. As DNA is synthesized in each cycle, more DNA is available to bind the dye, and so the fluorescence increases until the PCR reaction reaches the plateau phase. A limitation of the use of SYBR Green I is that any PCR product that is amplified is capable of binding the dye, resulting in increased fluorescence. The PCR primers

Fig. 17. The TaqMan principle. During PCR, a TaqMan probe hybridizes to one of the strands that is being amplified. The probe contains a fluorescent reporter (F) and a quencher (Q). The quencher prevents fluorescence by the reporter when the two molecules are physically in close proximity. Taq polymerase extends a primer during PCR, and the enzyme's 5'-3' exonuclease activity degrades the TaqMan probe. This separates the fluorescent reporter from the quencher, allowing the former to fluoresce. Increases in fluorescence therefore indicate that the sequence to which the TaqMan probe hybridizes are present in the sample and are being amplified by PCR. Measurements of increases in fluorescence during PCR can be used for quantitative PCR assays.

Fig. 17. The TaqMan principle. During PCR, a TaqMan probe hybridizes to one of the strands that is being amplified. The probe contains a fluorescent reporter (F) and a quencher (Q). The quencher prevents fluorescence by the reporter when the two molecules are physically in close proximity. Taq polymerase extends a primer during PCR, and the enzyme's 5'-3' exonuclease activity degrades the TaqMan probe. This separates the fluorescent reporter from the quencher, allowing the former to fluoresce. Increases in fluorescence therefore indicate that the sequence to which the TaqMan probe hybridizes are present in the sample and are being amplified by PCR. Measurements of increases in fluorescence during PCR can be used for quantitative PCR assays.

and reaction conditions must be chosen so as to ensure specificity of the reaction if reliable quantification is to be achieved. The second real-time PCR monitoring approach involves the use of target-specific probes, namely, TaqMan probes and molecular beacons.

TaqMan. Real-time PCR can be used to quantify a specific DNA PCR product by use of FRET probes (17). This form of realtime PCR increases the specificity of the reaction considerably over that achieved by use of SYBR Green I. The principle of FRET technology is shown in Fig. 17. A FRET probe is an oligonucleotide that has a fluorophore at its 5' end and a fluorescence quencher at its 3' end. The distance between these is such that when the fluorophore is excited, its energy is absorbed by the quencher, and so the overall fluorescence of the reaction is low. The middle region of the

Molecular Beacon

Molecular Beacon

Amplified Nucleic Acid

Fig. 18. The molecular beacon principle. A molecular beacon is an oligonucleotide that contains a fluorescent reporter (F) and a quencher (Q) at its ends. (A) Because of sequence complementarity near the ends, the structure forms a hairpin. The central part is complementary to a target DNA that is amplified by PCR. (B) If the target DNA is present, the molecular beacon hybridizes to it, thus separating the reporter and quencher. The former is now able to fluoresce.

oligonucleotide is designed to hybridize specifically to the region of DNA being amplified. As the target is replicated, the TaqMan probe hybridizes to its complementary sequence. As the PCR primer is extended by Taq polymerase to the hybridized probe, the polymerase removes the FRET probe by 5'-3' nuclease cleavage. This releases the free fluorophore into solution, where its fluorescence is unimpeded by the quencher. The fluorescence of the reaction therefore increases, and this increase is in proportion to the amount of amplified product present. This allows for real-time monitoring of the amplification of a specific DNA target. Because the rate of amplification is dependent on the amount of template at the beginning of PCR, the reaction can be used to quantify a nucleic acid target. Coamplification of an internal control can be monitored by use of a specific TaqMan probe.

Molecular Beacons. A molecular beacon, like a TaqMan probe, is an oligonucleotide that contains a fluorophore and a quencher (Fig. 18). These are at the ends of a stem-loop structure. The loop is complementary to a sequence in a target nucleic acid, and the stem contains a short stretch of complementary sequences such as the structure

Fig. 19. Invader principle. (A) A primary probe is hybridizing to its target sequence in DNA. The Invader probe overlaps by 1 bp. This arrangement is recognized by Cleavase, which cuts the primary probe, liberating the "5'-flap." (B) The liberated flap functions as an Invader oligonucleotide in a second reaction with a FRET cassette. Cleavase cuts the FRET cassette, leading to separation of the fluorescent reporter (F) from the quencher (Q). (C) The former can now fluoresce.

Fig. 19. Invader principle. (A) A primary probe is hybridizing to its target sequence in DNA. The Invader probe overlaps by 1 bp. This arrangement is recognized by Cleavase, which cuts the primary probe, liberating the "5'-flap." (B) The liberated flap functions as an Invader oligonucleotide in a second reaction with a FRET cassette. Cleavase cuts the FRET cassette, leading to separation of the fluorescent reporter (F) from the quencher (Q). (C) The former can now fluoresce.

folds as shown. The fluorophore and the quencher are physically adjacent when the structure is folded, and this inhibits fluorescence. When the target nucleic acid is present, the loop can hybridize to its complementary sequences. This results in disruption of the stem-loop structure and physical separation of the quencher from the fluorophore, which results in an increase in fluorescence of the solution.

Invader®. Invader is a homogenous, isothermal mutation detection system produced by Third Wave Technologies (Madison, WI) (18). It is used to identify single-nucleotide polymorphisms and point mutations in genomic DNA. The principle of Invader is shown in Figure 19. An Invader reaction uses two probes, a primary probe and an Invader probe. The primary probe contains a 3' region that can anneal to the target region in genomic DNA and a 5' region that is not complementary to the target and therefore does not hybridize to the target. The latter region of the primary probe is termed the flap. At the junction between these two regions is the base that is complementary to the base in the target DNA that is to be identified as being present or absent. In Fig. 19, this is a C in the target DNA, and the primary probe contains the complementary G. The Invader probe is complementary to the target DNA 5' of the base to be tested and extends in a 3' direction to overlap that base. If this base is present in the target DNA, the primary probe is cut by an enzyme known as Cleavase. This liberates the flap region from the primary probe. The temperature at which the Invader reaction takes place is close to the Tm of the primary probe, and the residual portion of this probe dissociates from the template and is replaced by another intact primary probe. The Tm of the Invader probe is designed to be at least 10°C higher than that of the primary probe, and therefore the Invader probe remains bound to the target as a series of primary probes binds and are cleaved.

The liberated flap then participates in a second Invader reaction that involves binding to an oligonucleotide containing a fluorophore and a quencher. This structure is known as a FRET cassette. When the fluorophore and the quencher molecules are physically adjacent, as in the FRET cassette, fluorescence is diminished. Binding of the flap allows for Cleavase to cut the FRET cassette, thereby separating the fluorophore from the quencher and increasing fluorescence in the reaction well. If the target DNA does not include the base of interest, the primary probe is not cut by Cleavase, and therefore there is no increase in fluorescence.

For each point mutation or SNP to be tested by Invader, two reaction wells are used: one for the wild-type allele and one for the mutant allele or SNP. Each of these uses a unique, allele-specific primary probe. It is not essential that the Invader oligonucleotide be complementary to the base to be tested, and therefore a common Invader probe can be used for both reactions. The ratio of fluorescence produced in both wells is used to determine the genotype.

Approaches to Detect Unknown Mutations

In certain molecular diagnostic assays, the goal is to screen a gene for any of multiple possible mutations. An efficient approach to achieve this goal is to apply a mutation screening technique that

Fig. 20. Single-strand conformation polymorphism (SSCP; left) and heteroduplex principles (right). (Left) Two samples of DNA, which differ in their sequence by just one nucleotide, are amplified by PCR. During the amplification, these are labeled, e.g., by incorporation of a radioisotope. Following PCR, the fragments are made single stranded and electrophoresed. Differences in sequence lead to different structures that exhibit differences in electrophoretic mobility in a nonde-naturing gel. This enables sequence variants to be identified using this technique. Characterization of the actual sequence variation requires further analysis. (Right) A known wild-type control sample and a sample containing a mutation are both amplified by PCR. Following amplification the two products are heated to melt the strands and then allowed to cool. This leads to formation of some het-eroduplexes (double-stranded DNA molecules that do not have perfectly complementary sequences). Heteroduplexes migrate more slowly than homoduplexes during electrophoresis. The presence of a heteroduplex band indicates a sequence variant in the sample being tested. Characterization of the actual sequence variation requires further analysis. WT is wild type, M is mutant.

can rapidly identify samples containing a sequence variation, and subsequently to confirm suspected mutations with a definitive method such as sequencing. Two common methods of screening for unknown mutations are single-strand conformation polymorphism (SSCP) analysis and heteroduplex analysis. The principles of these methods are shown in Fig. 20.

Single-Strand Conformational Polymorphism Analysis

In dilute solutions, single-stranded DNA molecules spontaneously adopt a three-dimensional conformation because of the formation of sequence-specific intramolecular base-pairing. Minor alterations to the sequence of the DNA strand can result in different conformations. These conformational differences alter the electrophoretic mobility of the DNA strand in nondenaturing gels. This principle is used in SSCP analysis. In a typical analysis, a region of DNA to be examined for the presence of mutations is amplified by PCR, and the PCR products are denatured (i.e., made single-stranded) and then electrophoresed in a nondenaturing gel. Control samples containing either the wild-type sequence or known mutations are run in parallel with patient samples. Electrophoretic mobility differences between the wild-type control and a patient sample indicate the presence of a possible mutation. The mutation is then characterized by sequencing.

Heteroduplex Analysis

A heteroduplex is a molecule composed of two strands of different sequence. Normally, DNA forms a double-stranded (duplex) molecule with two strands that are complementary to each other. However, a minor degree of noncomplementarity between the paired strands can be tolerated in a molecule that can still exist as a duplex. Such heteroduplexes migrate more slowly during electrophoresis than does a homoduplex (in which there is complete complementarity between strands) of similar size. Heteroduplexes are generated by mixing amplified mutant DNA with amplified wild-type DNA, heating the mixture to separate the strands, and then cooling to allow duplexes to form. Both homoduplexes and heteroduplexes will be produced on cooling. When the products from this procedure are separated by electrophoresis, a heteroduplex band will be visible as a more slowly migrating band than that formed by homoduplex molecules.

Denaturing and Temperature Gradient Gel Electrophoresis

The concentration of denaturing agents, or temperature, at which the two strands in a DNA molecule separate ("melt") is determined by the sequence of the DNA. If a DNA molecule is electrophoresed into an environment with an increasing concentration of a denaturing agent, or into a gradient of increasing temperature, small sections of the molecule with the weakest interstrand bonding will begin to separate. This alteration will slow the electrophoretic mobility of the molecule. If molecules with sequence variations are run in parallel lanes in such a gel, these will show different electrophoretic mobili ties. The use of progressively denaturing conditions can also be used to increase the separation between homoduplex and heteroduplex molecules in a heteroduplex analysis.

DNA Sequencing

DNA sequencing is considered the gold standard for mutation identification. However, the limited sample throughput of many automated DNA sequencers has tended to preclude its routine application in many clinical laboratories. The most common method of DNA sequencing is based on the chain termination method reported by Sanger, who won a Nobel prize in 1980 for this technique (19).

In the chain termination method for DNA sequencing, the piece of DNA to be sequenced is first amplified by PCR or other cloning strategy. The amplified DNA is denatured, and a primer is allowed to anneal to one end of the DNA. This primer functions in essentially the same way as a primer that might be used for PCR, namely, it serves as an anchor and starting point for a DNA polymerase to extend the primer using the sample DNA as a template. Only one primer is used for each sequencing reaction, which means that one of the two strands of DNA is extended. The sequencing reaction includes both dNTPs and one of the four di-deoxynucleotide triphosphates (ddNTPs). The latter do not have the 3'-hydroxyl group that is present in dNTPs. Once a dideoxynucleotide is incorporated in a molecule of DNA, no further extension is possible because there is no available hydroxyl group on the sugar group for the formation of the 3'-5' phosphodiester linkage that is found in nucleic acid polymers. At each position in the template at which the complementary base to the ddNTP is present, the polymerase can incorporate either the ddNTP or the normal dNTP. The relative likelihood of incorporating either nucleotide is dependent on the relative concentrations of ddNTP and dNTP. If the concentration of ddNTP is relatively high, DNA synthesis will tend to terminate early and produce large numbers of shorter molecules. Conversely, if the concentration of ddNTP is relatively low, longer length fragments can be synthesized before synthesis is terminated by a dideoxynucleotide. By controlling the concentrations of dNTP and ddNTP, it is possible to generate a range of fragment sizes.

A sequence analysis can be performed by setting up four separate reactions, each of which contains one of the ddNTPs (i.e., ddATP,

Fig. 21. Chain termination sequencing principle. A series of primer extension reactions is performed using a DNA template to be sequenced (top). The reactions contain mixtures of the deoxynucleotide triphosphates and di-deoxynucleotide triphosphates. Each reaction is terminated when a di-deoxynucleotide is incorporated. Separation of the products of these reactions in polyacrylamide gels enables determination of the sequence of the DNA template by reading the size order of the products (lower left). If each of the four di-deoxynucleotides is labeled with a different fluorescent dye, an automated reading of the products during size separation (e.g., by capillary electrophoresis) can be performed (lower right).

Fig. 21. Chain termination sequencing principle. A series of primer extension reactions is performed using a DNA template to be sequenced (top). The reactions contain mixtures of the deoxynucleotide triphosphates and di-deoxynucleotide triphosphates. Each reaction is terminated when a di-deoxynucleotide is incorporated. Separation of the products of these reactions in polyacrylamide gels enables determination of the sequence of the DNA template by reading the size order of the products (lower left). If each of the four di-deoxynucleotides is labeled with a different fluorescent dye, an automated reading of the products during size separation (e.g., by capillary electrophoresis) can be performed (lower right).

ddCTP, ddGTP, or ddTTP) (Fig. 21). These reactions will generate a range of fragment lengths, each of which terminate in the respective ddNTP. By electrophoretic separation of the products of these reactions in adjacent lanes, it is possible to determine the sequence of one strand of a DNA molecule. An alternative approach, which utilizes only one sequencing reaction, involves the use of chain termination nucleotides that are linked to dyes with different fluorescence

_RNA Sample

_ DNA Template

^ with T7 promoter

In Vitro Translation with 35S-Methionine

Radiolabeled Peptides

Gel Electrophoresis of Peptides

Fig. 22. Protein truncation assay principle. An RNA from the gene of interest is amplified by RT-PCR, incorporating a T7 RNA polymerase promoter. This product is used for in vitro transcription and translation. The peptides produced are labeled by incorporation of 35S-methionine. The size of these peptides is determined by electrophoresis in denaturing gels. A shortened peptide indicates the presence of a truncating mutation in the RNA.

wavelengths. The products of such a reaction are separated by a technique such as capillary electrophoresis. As the separated molecules pass through the detection window, they are excited by UV light, and the identity of the passing fluorophore (and therefore the base) is determined from its emitted light spectrum.

Protein Truncation Test

The protein truncation test is a method of identifying nonsense mutations in genes (20) (Fig. 22). These give rise to premature stop codons that cause early termination of protein synthesis. By performing an in vitro transcription and translation using a cloned gene or cDNA, the length of the translated protein can be determined. The cloned construct contains a T7 RNA polymerase promoter at its 5' end and a eukaryotic translation initiation codon (AUG). This is used as a template by T7 RNA polymerase to produce RNA. In vitro translation in a suitable system (e.g., a reticulocyte lysate) generates peptides, and the length of the translated peptides can be determined

T7 Promoter

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Fig. 23. Four-part photomicrograph of HER2/neu gene amplification detection by FISH. (Upper left) Unamplified case with a mean signal count of 1.5 signals per nucleus. (Upper right) Breast cancer with a borderline result featuring a mean signal count of 4.1 signals per nucleus. (Lower left) Significantly amplified breast tumor with a mean signal count of 17.5 signals per nucleus. (Lower right) Another example of a significantly amplified breast cancer with a mean HER2/neu signal count of 24. 9 signals per nucleus (Reproduced with permission from ref. 1).

Fig. 23. Four-part photomicrograph of HER2/neu gene amplification detection by FISH. (Upper left) Unamplified case with a mean signal count of 1.5 signals per nucleus. (Upper right) Breast cancer with a borderline result featuring a mean signal count of 4.1 signals per nucleus. (Lower left) Significantly amplified breast tumor with a mean signal count of 17.5 signals per nucleus. (Lower right) Another example of a significantly amplified breast cancer with a mean HER2/neu signal count of 24. 9 signals per nucleus (Reproduced with permission from ref. 1).

by electrophoresis in a denaturing gel. The presence of a premature truncating mutation is indicated by synthesis of a short peptide. Because of its complexity, the protein truncation test is primarily used as a research tool. It can be used to identify truncating mutations in genes in which this kind of mutation is common, e.g., BRCA1, BRCA2, or DMD (21,22).

Fluorescence In Situ Hybridization

Fluorescence in situ hybridization (FISH) is a cytogenetic technique in which a fluorescently labeled probe is hybridized to chromosomes. Probes used in FISH are either fluorescently labeled (direct technique) or contain a hapten such as biotin or digoxigenin that can be detected by a fluorescently labeled conjugate (indirect technique). Hybridization is detected by fluorescence microscopy (Fig. 23).

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Fig. 24. Four-color FISH detection of the t(9;22) BCR-ABL translocation. (A) A normal cell has been hybridized with four FISH probes. The blue and red probes hybridize to sequences in chromosome 22. The yellow and green probes hybridize to sequences in chromosome 9. The der(22) chromosome, which contains the BCR-ABL fusion, is shown by the adjacent green and red probes. The der(9) chromosome, which contains the reciprocal ABL-BCR fusion, is shown by the adjacent yellow and blue probes (Courtesy of Cancer Genetics, Inc.).

Fig. 24. Four-color FISH detection of the t(9;22) BCR-ABL translocation. (A) A normal cell has been hybridized with four FISH probes. The blue and red probes hybridize to sequences in chromosome 22. The yellow and green probes hybridize to sequences in chromosome 9. The der(22) chromosome, which contains the BCR-ABL fusion, is shown by the adjacent green and red probes. The der(9) chromosome, which contains the reciprocal ABL-BCR fusion, is shown by the adjacent yellow and blue probes (Courtesy of Cancer Genetics, Inc.).

FISH can be performed on metaphase chromosome preparations (i.e., from cells that can be cultured in vitro) or directly on nondividing cells (interphase FISH) (23). A large number of probes is commercially available that are used for different kinds of applications. For example, probes for a-satellite markers can be used to identify the origin of marker chromosomes. Probes for specific loci or chromosomal regions can be used to demonstrate deletions or amplifications (Fig. 23). FISH is used to demonstrate abnormalities in the subtelomeric regions of chromosomes that are responsible for a number of cases of unexplained mental retardation. By using several probes labeled with different fluo-rochromes, it is possible to detect chromosomal translocations. In this approach, the probes hybridize to their corresponding sequences on each of the partner chromosomes in the translocation and on the normal chromosomes. The presence of a translocation is indicated by the close proximity of probes that hybridize to the translocation partners (Fig. 24). Among other applications, interphase FISH is used for rapid identification of the common chromosomal aneuploidies in prenatal samples.

Spectral Karyotyping and Multiplex FISH

Spectral karyotyping (SKY) (24) and multiplex FISH (M-FISH) (25) are related analytical techniques that are developments of FISH. In both techniques, fluorescently labeled probes for each of the 24 chromosomes are applied to metaphase chromosome spreads and visualized using a computerized imaging system. The probes are produced by flow sorting or microdissecting normal chromosomes and then performing PCR on the isolated chromosomes using degenerate primers. For each chromosome, the PCR-generated probes are synthesized so as to contain a unique combination of different fluorophores. In this way, each chromosome is represented by probes that have unique fluorospectroscopic characteristics. In general, N fluorophores can be used to generate 2N - 1 unique combinations of labeled probes. Five fluorophores can be used to produce 31 possible combinations of dyes, which are sufficient to label each of the 24 chromosomes with a unique combination of fluorophores.

After hybridization, the labeled probes are bound to the metaphase chromosomes in the sample to be analyzed. The fluorescent probes are excited and the image is captured. By computer analysis of the image, the particular set of probes that is hybridizing to a metaphase chromosome can be identified. The results are displayed in false colors for ease of interpretation, with each chromosome being assigned a unique false color (Fig. 25).

SKY and M-FISH differ primarily in the method by which the image is analyzed. In SKY, an interferometer is used to identify the combination of fluorophores hybridized to each chromosome. In M-FISH, the image is photographed using a set of filters. Computer analysis of the image obtained with each filter is used to determine which probes have hybridized to each chromosome.

Uses and Limitations of SKY and M-FISH

Both techniques are of value for rapid detection of chromosomal aneuploidy and identification of the chromosome involved. Translocations, and the chromosomes involved in translocations, can also be rapidly identified. The origin of marker chromosomes can be established in some cases. However, these techniques are not suit-

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