Generation of DNA Fragments

Although fragments of DNA can be produced by mechanical shearing techniques, this is a haphazard process producing fragments that vary in size. In the early 1970s, it was recognized that certain microbes contain enzymes that cleave double-stranded DNA in or near a particular sequence of nucleotides. These enzymes restrict the entry of foreign DNA into bacterial cells and were therefore called restriction enzymes. They recognize a palindromic nucleotide sequence of DNA of between four and eight nucleotides in length (i.e., the same sequence of nucleotides occurring on the two complementary DNA strands when read in one direction of polarity, e.g., 5′ to 3′) (Table 4.2). The longer the nucleotide recognition sequence of the restriction enzyme, the less frequently that particular nucleotide sequence will occur by chance and therefore the larger the average size of the DNA fragments generated.

More than 300 different restriction enzymes have been isolated from various bacterial organisms. Restriction endonucleases are named according to the organism from which they are derived (e.g., EcoRI is from Escherichia coli and was the first restriction enzyme isolated from that organism).

The complementary pairing of bases in the DNA molecule means that cleavage of double-stranded DNA by a restriction endonuclease always creates double-stranded breaks, which, depending on the cleavage points of the particular restriction enzyme used, results in either a staggered or a blunt end (Figure 4.2).

FIGURE 4.2 The staggered and blunt ends generated by restriction digest of double-stranded DNA by EcoRI and SmaI. Sites of cleavage of the DNA strands are indicated by arrows.

Digestion of DNA from a specific source with a particular restriction enzyme will produce the same reproducible collection of DNA fragments each time the process is carried out.

Recombination of DNA Fragments

DNA from any source, when digested with the same restriction enzyme, will produce DNA fragments with identical complementary ends or termini. When DNA has been cleaved by a restriction enzyme that produces staggered termini, these are referred to as being ‘sticky’ or ‘cohesive’ because they will unite under appropriate conditions with complementary sequences produced by the same restriction enzyme on DNA from any source. Initially the cohesive termini are held together by hydrogen bonding but are covalently attached with the enzyme called DNA ligase. The union of two DNA fragments from different sources produces what is referred to as a recombinant DNA molecule.

Vectors

A vector is the term for the carrier DNA molecule used in the cloning process that, through its own independent replication within a host organism, will allow the production of multiple copies of itself. The incorporation of the target DNA into a vector allows the production of large amounts of that DNA fragment.

For naturally occurring vectors to be used for DNA cloning, they need to be modified to ensure that the target DNA is inserted at a specific location and that recombinant vectors containing target inserted DNA can be detected. Many of the early vectors were constructed so that insertion of the target DNA in a gene for antibiotic resistance resulted in loss of that function (Figure 4.3).

FIGURE 4.3 Two plasmids originally used in recombinant DNA technology showing drug resistance genes (Ap^r, ampicillin resistance; Tc^r, tetracycline resistance) and cleavage sites of restriction endonucleases that are present in the DNA only once for use as a cloning site.

The five main types of vector commonly used include plasmids, bacteriophages, cosmids, and bacterial and yeast artificial chromosomes (BACs and YACs). The choice of vector used in cloning depends on a number of factors, such as the particular restriction enzyme being used and the size of the target DNA to be inserted. Some of the early vectors, such as plasmids and bacteriophages, were very limited in terms of the size of the target DNA fragment that could be inserted. Later generations of vectors, such as cosmids, can take inserts up to approximately 50 kb in size. A cosmid is essentially a plasmid that has had all but the minimum vector DNA necessary for propagation removed (i.e., the cos sequence), to enable insertion of the largest possible foreign DNA fragment and still allow replication.

The development of BACs and YACs allows the possibility of cloning DNA fragments of between 300 kb and 1000 kb in size. YACs consist of a plasmid that contains within it the minimum DNA sequences necessary for centromere and telomere formation plus DNA sequences known as autonomous replication sequences, all of which are necessary for accurate replication within yeast. YACs have the advantage that they can incorporate DNA fragments of up to 1000 kb in size as well as allow replication of eukaryotic DNA with repetitive DNA sequences, which often cannot take place in bacterial cells. Many eukaryotic genes are very large, being up to 2 to 3 million base pairs (bp) in length (p. 388). YACs allow detailed mapping of genes of this size and their flanking regions, whereas the use of conventional vectors would require an inordinate number of overlapping clones.

Transformation of the Host Organism

After introducing the target DNA fragment into the vector, the recombinant vector is introduced into specially modified bacterial or yeast host cells. The bacterial cell membrane is not normally permeable to large molecules such as DNA fragments but can be made permeable by a variety of different methods, including exposure to certain salts or high voltage; this is known as becoming competent. Usually only a single DNA molecule is taken up by a host cell undergoing the process known as transformation. If the transformed cells are allowed to multiply, large quantities of identical copies of the original single target DNA or clones will be produced (Figure 4.4).

FIGURE 4.4 Generation of a recombinant plasmid using EcoRI and transformation of the host bacterial organism.

(From Emery AE 1981 Recombinant DNA technology. Lancet ii:1406–1409, with permission.)

Screening for Recombinant Vectors

After the transformed cells have multiplied in culture medium, they are plated out on a master plate of nutrient agar in a Petri dish. Recombinant vectors can be screened for by a detection system; for example, loss of antibiotic resistance can be screened for by replica plating on agar containing the appropriate antibiotic (see Figure 4.3). Thus, if the enzyme PstI were used to generate DNA fragments and to cut the plasmid pBR322, any recombinant plasmids produced would make the bacterial host cells they transform sensitive to ampicillin, as this gene would no longer be functional, but they would remain resistant to tetracycline. Replica plating of the master plates from the cultures allows identification of individual specific recombinant clones.

Selection of Specific Clones

Several techniques have been developed to detect the presence of clones with specific DNA sequence inserts. The most widely used method is nucleic acid hybridization (p. 57). Colonies of transformed host bacteria with recombinant clones are used to make replica plates that are lyzed and then blotted on to a nitrocellulose filter to which nucleic acid binds. The DNA of the replica blot is then denatured to make the DNA single stranded, which will allow it to hybridize with single-stranded, radioactively labeled DNA or RNA probes (p. 58), which can then be detected by exposure to an x-ray film, or what is known as autoradiography. In this way, a transformed host bacterial colony containing a sequence complementary to the probe can be detected and, from its position on the replica plate, the colony containing that clone can be identified on the master plate, ‘picked’, and cultured separately (Figure 4.5).

FIGURE 4.5 Identification of recombinant DNA clones with specific DNA inserts by loss of antibiotic resistance, nucleic acid hybridization, and autoradiography.

The PCR

DNA sequence information is used to design two oligonucleotide primers (amplimers) of approximately 20 bp in length complementary to the DNA sequences flanking the target DNA fragment. The first step is to denature the double-stranded DNA by heating. The primers then bind to the complementary DNA sequences of the single-stranded DNA templates. DNA polymerase extends the primer DNA in the presence of the deoxynucleotide triphosphates (dATP, dCTP, dGTP, and dTTP) to synthesize the complementary DNA sequence. Subsequent heat denaturation of the double-stranded DNA, followed by annealing of the same primer sequences to the resulting single-stranded DNA, will result in the synthesis of further copies of the target DNA. Some 30 to 35 successive repeated cycles results in more than 1 million copies (amplicons) of the DNA target, sufficient for direct visualization by ultraviolet fluorescence after ethidium bromide staining, without the need to use indirect detection techniques (Figure 4.6).

FIGURE 4.6 Diagram of the polymerase chain reaction showing serial denaturation of DNA, primer annealing, and extension with doubling of the target DNA fragment numbers in each cycle.

PCR allows analysis of DNA from any cellular source containing nuclei; in addition to blood, this can include less invasive samples such as buccal scrapings or pathological archival material. It is also possible to start with quantities of DNA as small as that from a single cell, as is the case in preimplantation genetic diagnosis (p. 335). Great care has to be taken with PCR, however, because DNA from a contaminating extraneous source, such as desquamated skin from a laboratory worker, will also be amplified. This can lead to false-positive results unless the appropriate control studies are used to detect this possible source of error.

Another advantage of PCR is the rapid turnaround time of samples for analysis. Use of the heat-stable Taq DNA polymerase isolated from the bacterium Thermophilus aquaticus, which grows naturally in hot springs, generates PCR products in a matter of hours rather than the days or weeks required for cell-based in-vivo DNA cloning techniques.

Real-time PCR machines have reduced this time to less than 1 hour, and fluorescence technology is used to monitor the generation of PCR products during each cycle, thus eliminating the need for gel electrophoresis.

DNA cloning by PCR, in contrast to in-vivo cell-based techniques, has the disadvantage that it requires knowledge of the nucleotide sequence of the target DNA fragment and is best used to amplify DNA fragments of up to 1 kb, although long-range PCR allows the amplification of larger DNA fragments of up to 20 kb to 30 kb.

Southern Blotting

Southern blotting, named after Edwin Southern (who developed the technique), involves digesting DNA by a restriction enzyme that is then subjected to electrophoresis on an agarose gel. This separates the DNA or restriction fragments by size, the smaller fragments migrating faster than the larger ones. The DNA fragments in the gel are then denaturated with alkali, making them single stranded. A ‘permanent’ copy of these single-stranded fragments is made by transferring them on to a nitrocellulose filter that binds the single-stranded DNA, the so-called Southern blot. A particular target DNA fragment of interest from the collection on the filter can be visualized by adding a single-stranded ³²P radioactively labeled DNA probe that will hybridize with homologous DNA fragments in the Southern blot, which can then be detected by autoradiography (Figure 4.7). Non-radioactive Southern blotting techniques have been developed with the DNA probe labeled with digoxigenin and detected by chemiluminescence. This approach is safer and generates results more rapidly. An example of the use of Southern blotting for diagnostic fragile X testing in patients is shown in Figure 4.8.

FIGURE 4.7 Diagram of the Southern blot technique showing size fractionation of the DNA fragments by gel electrophoresis, denaturation of the double-stranded DNA to become single stranded, and transfer to a nitrocellulose filter that is hybridized with a ³²P radioactively labeled DNA probe.

FIGURE 4.8 Southern blot to detect methylation of the FMR1 promoter in patients with fragile X. DNA digested with EcoR1 and the methylation sensitive enzyme Bst Z1 was probed with Ox1.9, which hybridizes to a CpG island within the FMR1 promoter. Patient 1 is a female with a methylated expansion, patients 2, 3, and 6 are normal females, patient 4 is an affected male and patient 5 is a normal male.

(Courtesy A. Gardner, Department of Molecular Genetics, Southmead Hospital, Bristol, UK.)

Northern Blotting

Northern blotting differs from Southern blotting by the use of mRNA as the target nucleic acid in the same procedure; mRNA is very unstable because of intrinsic cellular ribonucleases. Use of ribonuclease inhibitors allows isolation of mRNA that, if run on an electrophoretic gel, can be transferred to a filter. Hybridizing the blot with a DNA probe allows determination of the size and quantity of the mRNA transcript, a so-called Northern blot. With the advent of real-time reverse transcriptase PCR, and microarray technology for gene expression studies, Northern blotting is used less often.

DNA Microarrays

DNA microarrays are based on the same principle of hybridization but on a miniaturized scale, which allows simultaneous analysis of several million targets. Short, fluorescently labeled oligonucleotides attached to a glass microscope slide can be used to detect hybridization of target DNA under appropriate conditions. The color pattern of the microarray is then analyzed automatically by computer. Four classes of application have been described: (1) expression studies to look at the differential expression of thousands of genes at the mRNA level; (2) analysis of DNA variation for mutation detection and single nucleotide polymorphism (SNP) typing (p. 67); (3) testing for genomic gains and losses by array comparative genomic hybridization (CGH) (p. 36); and (4) a combination of the latter two, SNP–CGH, which allows the detection of copy-neutral genetic anomalies such as uniparental disomy (p. 121).

Size Analysis of PCR Products

Deletion or insertion mutations can sometimes be detected simply by determining the size of a PCR product. For example, the most common mutation that causes cystic fibrosis, p.Phe508del, is a 3-bp deletion that can be detected on a polyacrylamide gel. Some trinucleotide repeat expansion mutations can be amplified by PCR (Figure 4.9).

FIGURE 4.9 Amplification of the GAA repeat expansion mutation by polymerase chain reaction (PCR) to test for Friedreich ataxia. Products are stained with ethidium bromide and electrophoresed on a 1.5% agarose gel. Lanes 1 and 8 show 500-bp ladder-size standards, lanes 2 and 4 show patients with homozygous expansions, lanes 3 and 6 show unaffected controls, lane 5 shows a heterozygous expansion carrier, and lane 7 is the negative control.

(Courtesy K. Thomson, Department of Molecular Genetics, Royal Devon and Exeter Hospital, Exeter, UK.)

Restriction Fragment Length Polymorphism

If a base substitution creates or abolishes the recognition site of a restriction enzyme, it is possible to test for the mutation by digesting a PCR product with the appropriate enzyme and separating the products by electrophoresis (Figure 4.10).

FIGURE 4.10 Detection of the HFE gene mutation C282Y by restriction fragment length polymorphisms (RFLP). The normal 387-bp polymerase chain reaction (PCR) product is digested with RsaI to give products of 247 and 140 bp. The C282Y mutation creates an additional recognition site for RsaI, giving products of 247, 111, and 29 bp. Lane 1 shows a 100-bp ladder-size standard. Lanes 2 though 4 show patients homozygous, heterozygous, and normal for the C282Y mutation, respectively. Lane 5 is the negative control.

(Courtesy N. Goodman, Department of Molecular Genetics, Royal Devon and Exeter Hospital, Exeter, UK.)

Amplification-Refractory Mutation System (ARMS) PCR

Allele-specific PCR uses primers specific for the normal and mutant sequences. The most common design is a two-tube assay with normal and mutant primers in separate reactions together with control primers to ensure that the PCR reaction has worked. An example of a multiplex ARMS assay to detect 12 different cystic fibrosis mutations is shown in Figure 4.11.

FIGURE 4.11 Detection of CFTR mutations by two-tube amplification-refractory mutation system (ARMS)-polymerase chain reaction (PCR). Patient 1 is heterozygous for ΔF508 (p.Phe508del). Patient 2 is a compound heterozygote for p.Phe508del and c.1717-1G > A. Patient 3 is homozygous normal for the 12 mutations tested. Primers for two internal controls (ApoB and ODC) are included in each tube.

Oligonucleotide Ligation Assay

A pair of oligonucleotides is designed to anneal to adjacent sequences within a PCR product. If the pair is perfectly hybridized, they can be joined by DNA ligase. Oligonucleotides complementary to the normal and mutant sequences are differentially labeled and the products identified by computer software (Figure 4.12).

FIGURE 4.12 Detection of CFTR mutations using an oligonucleotide ligation assay. Multiplex polymerase chain reaction (PCR) amplifies 15 exons of the CFTR gene. Oligonucleotides are designed to anneal to the PCR products such that two oligonucleotides anneal to adjacent sequences for each mutation and are then joined by ligation. The 32 mutations are discriminated using a combination of size and differently colored fluorescent labels. This patient is a compound heterozygote for the ΔF508 (p.Phe508del) and c.1585-1G > A mutations.

(Courtesy Karen Stals, Department of Molecular Genetics, Royal Devon and Exeter Hospital, Exeter, UK.)

Real-Time PCR

There are multiple hardware platforms for real-time PCR and ‘fast’ versions that can complete a PCR reaction in less than 30 minutes. TaqMan^TM and LightCycler^TM use fluorescence technology to detect mutations by allelic discrimination of PCR products. Figure 4.13 illustrates the factor V Leiden mutation detected by TaqMan^TM methodology.

FIGURE 4.13 Real-time polymerase chain reaction (PCR) to detect the Factor V Leiden mutation. A, TaqMan technique. The sequence encompassing the mutation is amplified by PCR primers, P1 and P2. A probe, P3, specific to the mutation is labelled with two fluorophores. A reporter fluorophore, R, is attached to the 5′ end of the probe and a quencher fluorophore, Q, is attached to the 3′ end. During the PCR reaction, the 5′ exonuclease activity of the polymerase enzyme progressively degrades the probe, separating the reporter and quencher dyes, which results in fluorescent signal from the reporter fluorophore. B, TaqMan genotyping plot. Each sample is analysed with two probes, one specific for the wild-type and one for the mutation. The strength of fluorescence from each probe is plotted on a graph (wild-type on X-axis, mutant on Y-axis). Each sample is represented by a single point. The samples fall into 3 clusters representing the possible genotypes; homozygous wild-type, homozygous mutant or heterozygous.

(Courtesy Dr. E. Young, Department of Molecular Genetics, Royal Devon and Exeter Hospital, Exeter, UK.)

DNA Microarrays (DNA ‘Chips’)

DNA microarrays hold the promise of rapid mutation testing. They involve synthesizing custom-designed 20 bp to 25 bp oligonucleotide sequences for both the normal DNA sequence and known and/or possible single nucleotide substitutions of a gene. These are attached to a ‘chip’ in a structured arrangement in what is known as a microarray. The sample DNA being screened for a mutation is amplified by PCR, fluorescently labeled, and hybridized with the oligonucleotides in the microarray (Figure 4.14). Computer analysis of the color pattern of the microarray generated after hybridization allows rapid automated mutation testing. The prospect of gene-specific DNA chip microarrays may lead to a revolution in the speed and reliability of mutation screening, provided the technology is affordable and the technique can be demonstrated to be robust. The detection of known base substitutions and SNPs has been very successful, but screening for insertion mutations is more limited.

FIGURE 4.14 Detection of HNF1A mutations using a DNA microarray. The ‘HNF1A chip’ contains normal and mutant probes for 75 different mutations spotted in triplicate. Patient DNA was amplified by multiplex polymerase chain reaction (PCR) to yield fluorescently labeled products that were hybridized to the chip. A, Control sample. B, Patient heterozygous for an HNF1A mutation.

(Courtesy N. Huh, Samsung Advanced Institute of Technology, South Korea.)

Conformation-Sensitive Capillary Electrophoresis

Conformation-sensitive capillary electrophoresis is used to detect the presence of heteroduplexes using fluorescence technology. PCR products can be multiplexed by using multiple fluorescent dyes. An alteration in the DNA sequence can result in a different conformation, which has a different electrophoretic mobility, and an appropriate polymer can be used for identification.

High-Resolution Melt Curve Analysis

This technique employs a class of fluorescent dyes that intercalate with double-stranded, but not single-stranded, DNA. The intercalating dye is incorporated in the PCR reaction and the products are then heated to separate the two strands. Fluorescence levels decrease as the DNA strands dissociate and this ‘melting’ profile depends on the PCR product size and sequence (Figure 4.15). High-resolution melt curve analysis appears to be very sensitive and can be used for high-throughput mutation screening.

FIGURE 4.15 High-resolution melt curve analysis (HRM). Melting profiles for normal and mutant samples are shown after normalization to a control sample. Each variant has a different melting profile.

Sanger Sequencing

The ‘gold standard’ method of mutation screening is DNA sequencing using the dideoxy chain termination method developed in the 1970s by Fred Sanger. This method originally employed radioactive labeling with manual interpretation of data. The use of fluorescent labels detected by computerized laser systems has improved ease of use and increased throughput and accuracy. Today’s capillary sequencers can sequence around 1 Mb (1 million bases) per day.

Dideoxy sequencing involves using a single-stranded DNA template (e.g., denatured PCR products) to synthesize new complementary strands using a DNA polymerase and an appropriate oligonucleotide primer. In addition to the four normal deoxynucleotides, a proportion of each of the four respective dideoxynucleotides is included, each labeled with a different fluorescent dye. The dideoxynucleotides lack a hydroxyl group at the 3′ carbon position; this prevents phosphodiester bonding, resulting in each reaction container consisting of a mixture of DNA fragments of different lengths that terminate in their respective dideoxynucleotide, owing to chain termination occurring at random in each reaction mixture at the respective nucleotide. When the reaction products are separated by capillary electrophoresis, a ladder of DNA sequences of differing lengths is produced. The DNA sequence complementary to the single-stranded DNA template is generated by the computer software and the position of a mutation may be highlighted with an appropriate software package (Figure 4.16).

FIGURE 4.16 Fluorescent dideoxy DNA sequencing. The sequencing primer (shown in red) binds to the template and primes synthesis of a complementary DNA strand in the direction indicated (A). The sequencing reaction includes four dNTPs and four ddNTPs, each labeled with a different fluorescent dye. Competition between the dNTPs and ddNTPs results in the production of a collection of fragments (B), which are then separated by electrophoresis to generate an electropherogram (C). A heterozygous mutation, p.Gly44Cys (GGC > TGC; glycine > cysteine), is identified by the software.

Pyrosequencing

Pyrosequencing uses a sequencing by synthesis approach in which modified nucleotides are added and removed one at a time, with chemiluminescent signals produced after the addition of each nucleotide. This technology generates quantitative sequence data rapidly and an example of its application in the identification of KRAS mutations in patients with colorectal cancer is shown in Figure 4.17.

FIGURE 4.17 Detection of a KRAS mutation in a colorectal tumour by pyrosequencing. The upper panel shows a normal control, sequence A GGT CAA GAG G. In the lower panel is the tumour sample with the KRAS mutation p.Gln61Leu (c.182A > T).

(Courtesy Dr. L. Meredith, Institute of Medical Genetics, University Hospital of Wales, Cardiff.)

Next-Generation ‘Clonal’ Sequencing

The demand for low-cost sequencing has driven the development of high-throughput sequencing technologies that produce millions of sequences at once. Next (or second) generation ‘clonal’ sequencers use an in vitro cloning step to amplify individual DNA molecules by emulsion or bridge PCR (Figure 4.18). The cloned DNA molecules are then sequenced in parallel, either by pyrosequencing, by using reversible terminators or with a sequencing by ligation approach. A comparison with Sanger sequencing is shown in Table 4.5 and an example of a mutation identified by next generation sequencing is shown in Figure 4.19. So-called ‘third generation’ sequencers have recently been developed. They can generate massively parallel sequence data from single molecules due to their extremely sensitive lasers.

FIGURE 4.18 Next-generation ‘clonal’ sequencing. DNA is fragmented and adaptors ligated before clonal amplification on a bead or glass slide. Sequencing takes place in situ and incorporated bases are detected by direct light emission or scanning of fluorophores. Data analysis includes base calling and alignment to a reference sequence in order to identify mutations or polymorphisms.

(Courtesy Dr. R. Caswell, Peninsula Medical School, Exeter.)

Table 4.5 Sanger Sequencing Compared to Next-Generation ‘Clonal’ Sequencing

FIGURE 4.19 Detection of a TP53 mutation in a patient with Li Fraumeni syndrome. The reference sequence is shown at the top and the patient sequence below. A heterozygous C > T mutation (c.430C > T; p.Gln144X) is visible in 11 of the 25 reads shown.

(Courtesy Jo Morgan and Graham Taylor, Leeds Institute of Molecular Medicine, St. James’s Hospital, Leeds, UK.)

Single Nucleotide Polymorphisms

Around 1 in 1000 bases within the human genome shows variation. SNPs are most frequently biallelic and occur in coding and non-coding regions. If an SNP lies within the recognition sequence of a restriction enzyme, the DNA fragments produced by that restriction enzyme will be of different lengths in different people. This can be recognized by the altered mobility of the restriction fragments on gel electrophoresis, so-called restriction fragment length polymorphisms, or RFLPs. Early genetic mapping studies used Southern blotting to detect RFLPs, but current technology enables the detection of any SNP. DNA microarrays have led to the creation of a dense SNP map of the human genome and assist genome searches for linkage studies in mapping single-gene disorders (p. 293) and association studies in common diseases.

Variable Number Tandem Repeats

Variable number tandem repeats (VNTRs) are highly polymorphic and are due to the presence of variable numbers of tandem repeats of a short DNA sequence that have been shown to be inherited in a mendelian co-dominant fashion (p. 113). The advantage of using VNTRs over SNPs is the large number of alleles for each VNTR compared with SNPs, which are mostly biallelic.

Minisatellites

Alec Jeffreys identified a short 10-bp to 15-bp ‘core’ sequence with homology to many highly variable loci spread throughout the human genome (p. 17). Using a probe containing tandem repeats of this core sequence, a pattern of hypervariable DNA fragments could be identified. The multiple variable-size repeat sequences identified by the core sequence are known as minisatellites. These minisatellites are highly polymorphic, and a profile unique to an individual (unless they have an identical twin!) is described as a DNA fingerprint. The technique of DNA fingerprinting is used widely in paternity testing and for forensic purposes.

Microsatellites

The human genome contains some 50,000 to 100,000 blocks of a variable number of tandem repeats of the dinucleotide CA:GT, so-called CA repeats or microsatellites (p. 18). The difference in the number of CA repeats at any one site between individuals is highly polymorphic and these repeats have been shown to be inherited in a mendelian co-dominant manner. In addition, highly polymorphic trinucleotide and tetranucleotide repeats have been identified, and can be used in a similar way (Figure 4.23). These microsatellites can be analyzed by PCR and the use of fluorescent detection systems allows relatively high-throughput analysis. Consequently, microsatellite analysis has replaced DNA fingerprinting for paternity testing and establishing zygosity.

FIGURE 4.23 Analysis of a tetranucleotide microsatellite marker in a family with a dominant disorder. Genotyper software was used to label the peaks with the size of the polymerase chain reaction (PCR) products. The 200-bp allele is segregating with the disorder in the affected members of the family.

(Courtesy M. Owens, Department of Molecular Genetics, Royal Devon and Exeter Hospital, Exeter, UK.)

Clinical Applications of Gene Tracking

If a gene has been mapped by linkage studies but not identified, it is possible to use the linked markers to ‘track’ the mutant haplotype within a family. This approach may also be used for known genes where a familial mutation has not been found. Closely flanking or intragenic microsatellites are used most commonly, because of the lower likelihood of finding informative SNPs within families. Figure 4.24 illustrates a family in which gene tracking has been used to determine carrier risk in the absence of a known mutation. There are some pitfalls associated with this method: recombination between the microsatellite and the gene may give an incorrect risk estimate, and the possibility of genetic heterogeneity (where mutations in more than one gene cause a disease) should be borne in mind.

FIGURE 4.24 Gene tracking in a family with Duchenne muscular dystrophy where no mutation has been found in the affected proband, III.4. Analysis of markers A, B, and C has enabled the construction of haplotypes; the affected haplotype is shown by an orange box. Both of the proband’s sisters were at 50% prior risk of being carriers. Gene tracking shows that III.1 has inherited the low-risk haplotype and is unlikely to be a carrier, but III.3 has inherited the high-risk haplotype and is therefore likely to be a carrier of Duchenne muscular dystrophy. The risk of recombination should not be forgotten.

Infectious Disease

PCR can be used to detect the presence of DNA sequences specific to a particular infectious organism before conventional evidence such as an antibody response or the results of cultures is available. An example is the screening of blood products for the presence of DNA sequences from the human immunodeficiency virus (HIV) to ensure the safety of their use (e.g., screening pooled factor VIII concentrate for use in males with hemophilia A). Another example is the identification of DNA sequences specific to bacterial or viral organisms responsible for acute overwhelming infections, where early diagnosis allows prompt institution of the correct antibiotic or antiviral agent with the prospect of reducing morbidity and mortality. Real-time PCR techniques can generate rapid results, with some test results being available within 1 hour of a sample being taken. This methodology is particularly useful in the fight against methicillin-resistant Staphylococcus aureus (MRSA), as patients can be rapidly tested on admission to hospital. Anyone found to be MRSA-positive can be isolated to minimize the risk of infection to other patients.

Malignant Disease

PCR may assist in the diagnosis of lymphomas and leukemias by identifying translocations, for example t(9;22), which is characteristic of chronic myeloid leukemia (CML). The extreme sensitivity of PCR means that minimal residual disease may be detected after treatment for these disorders, and early indication of impending relapse will inform treatment options. For example, all patients with CML treated with the tyrosine kinase inhibitor Imatinib are regularly monitored as resistant clones may develop. After bone marrow transplantation, microsatellite markers may be used to monitor the success of engraftment by analysis of donor- and patient-specific alleles.