DNA at the Molecular Level

DNA is a duplex molecule composed of two complementary hydrogen-bonded nucleotide strands (Figure 32-4). Deoxyribonucleotides and ribonucleotides are the building blocks of DNA and RNA, respectively. Each nucleotide is composed of a 5-carbon sugar (pentose), a nitrogenous base, and a phosphate group (Figure 32-5). The numbers one prime (1′) to five prime (5′) designate the pentose’s carbons. In DNA, the pentose is a ribose in which the hydroxyl group on the 2′ carbon is replaced by a hydrogen molecule, hence 2′-deoxyribose. In RNA, the 2′ ribose retains the 2′ hydroxyl group. The hydroxyl group present on the 3′ carbon of the sugar is crucial for polymerization of the nucleotide monomers to form the nucleic acid strand.

The nitrogenous base is linked to the sugar by a glycosidic bond at the 1′ carbon. Four different bases form DNA, but the linkage to the sugar is the same for each. The phosphate group is linked to sugar at the 5′ carbon by a phosphodiester bond. The phosphate group is also crucial for addition of nucleotides to the growing polymer. A sugar, whether ribose or deoxyribose, linked to a nitrogenous base, but without a phosphate group, is called a nucleoside. A nucleoside cannot be incorporated into DNA, and neither can a nucleotide consisting of only one phosphate group (deoxynucleotide monophosphate, or dNMP). To be incorporated into a growing strand of DNA, the nucleotide must have three phosphate groups linked to one another, referred to as the α-, β-, and γ-phosphates. The α-phosphate is linked to the sugar.

Creation of a phosphodiester bond between the 3′ hydroxyl group of the existing strand and the 5′ α-phosphate of the nucleotide monomer requires the protein enzyme DNA polymerase. This enzyme recognizes the hydroxyl group on the 3′ carbon of the sugar and bonds the 3′ hydroxyl group of one nucleotide with the α-phosphate group of another (Figure 32-6). Polymerization of subsequent nucleotides forms a DNA strand.

DNA consists of two strands that are antiparallel and complementary (Figure 32-7). One strand begins with a phosphate group attached to the 5′ carbon of the first nucleotide oriented to the left and ends with the hydroxyl group on the 3′ carbon of the last nucleotide oriented to the right. This strand is in the 5′-to-3′ direction. The other strand runs in the 3′-to-5′ direction, or antiparallel. The nucleotide sequences composing these strands provide the encoded messages of our genes. Therefore, the addition of nucleotides is highly regulated.

One regulation mechanism arises from the complementary characteristic of the nucleotides. A nucleotide’s identity depends on the type of nitrogenous base present on the template. There are two categories of nitrogenous bases in nucleic acids, purines and pyrimidines (Figure 32-8). The bases adenine (A) and guanine (G) are double-ringed purines, whereas thymine (T) and cytosine (C) are single-ringed pyrimidines. Adenine forms hydrogen bonds at two points with thymine (A:T), whereas guanine forms hydrogen bonds at three points with cytosine (G:C). If a strand has a 5′-CTAG-3′ sequence, the complementary nucleotides on the 3′-to-5′ strand are 3′-GATC-5′. In RNA, the pyrimidine uracil (U) takes the place of thymine and forms hydrogen bonds with adenine. Hydrogen bonds between A:T and G:C hold the strands together (Figure 32-9). RNA is most often single-stranded.

In addition to conferring identity to the nucleotide, the nitrogenous bases assist in maintaining a constant width between the strands of a DNA molecule. DNA resembles a ladder, with the repeating sugar and phosphate groups forming the sides of the ladder and the bases forming the rungs. The pairing of a double-ringed purine on one strand with a single-ringed pyrimidine on the other maintains a consistent distance between the DNA strands. This makes DNA flexible, which allows the molecule to twist into a helix. Twisting stabilizes the molecule and protects the bases from their environment.

Transcription and Translation

DNA provides a permanent set of instructions. The cellular enzyme RNA polymerase transcribes the code. RNA polymerase recognizes starter sequences called promoters. Promoters lie upstream of coding sequences and bind RNA polymerase to separating DNA strands. The enzyme then slides along the DNA strand 3′ to 5′, “reading” the code and polymerizing (assembling) the complementary ribonucleotides. As the complementary ribonucleotides form hydrogen bonds with the bases of the exposed DNA strand, the RNA polymerase creates phosphodiester bonds to extend the single-stranded primary RNA transcript (Figure 32-10). If the nucleotide sequence of the DNA strand is 3′-CTAG-5′, the primary RNA transcript is 5′-GAUC-3′.

Primary mRNA segments are composed of introns and exons. Introns are untranslated intervening sequences located within the coding portions of genes. Their functions remain unclear, although they may play a role in regulation of gene expression.⁵ Exons are the sequences that encode the gene product. Before mRNA can serve as a translation template, the introns must be excised from the primary transcript and the exons adjoined. The mature mRNA is completed by the addition of a 5′ cap and a tail of many repeated adenine nucleotides.⁶ The mRNA leaves the nucleus and enters cytoplasmic ribosomes to be translated.

Ribosomes translate the mRNA code into a peptide sequence. Complexes of proteins and structural ribosomal RNAs (rRNAs) form both large and small ribosome subunits. Mature cytoplasmic mRNA is bound by the small ribosomal subunit at the translation start site. At this point, another series of elements is introduced, transfer RNAs (tRNAs), each bound to its specific amino acid. Because there are 20 natural amino acids, there are 20 tRNAs. Each tRNA has a specific nucleic acid sequence located at the point of interaction with the mRNA, complementary to the nucleotide sequence of the mRNA. Each tRNA interacting sequence (anticodon) complements a specific three-nucleotide sequence (codon) of the mRNA.

The mRNA codon AUG is the most common translation start site and codes for the amino acid methionine. The first step in translation is hydrogen bonding of the appropriately charged tRNA (with a bound methionine) to the start codon of the mRNA. The appropriate tRNA is then bonded to the adjacent codon and a peptide bond is catalyzed between the two amino acids. The peptide bond forms between the carboxyl terminus of the methionine in the existing peptide chain and the amino terminus of the amino acid to be added. Hydrogen bonding of tRNAs to the codons and the formation of the peptide bonds are mediated by the ribosome. With addition of more amino acids, translation proceeds until a termination site (nonsense codon) is reached. Three codons exist that do not code for any amino acid: UAA, UAG, and UGA. These terminate translation. The ribosome then dissociates, and the peptide folds to its functional shape.

DNA Replication and the Cell Cycle

After cells carry out their functions, they either divide via mitosis or die via apoptosis, also called programmed cell death. The cell cycle progresses through a sequence (Figure 32-11). Interphase is made up of the G₁, S, and G₂ phases. During the G₁ phase, the cell grows rapidly and performs its cellular functions. S phase is the synthesis stage, in which DNA is replicated. The G₂ phase is the period when the cell produces materials essential for cell division. The M phase refers to mitosis, during which two identical daughter cells are produced, each of which receives one entire set of the DNA that was replicated during S phase. Some cells exit the cell cycle during the G₁ phase and enter a phase called G₀. Cells in G₀ normally do not reenter the cell cycle and remain alive performing their function until apoptosis occurs.

DNA replication during the S phase requires a complex orchestration of events; this discussion focuses on those events that are exploited for molecular diagnostic testing. Contained within the double-stranded DNA helix are multiple origins of replication. At each origin, the enzyme helicase disrupts and untwists the hydrogen bonds, separating the DNA strands and producing two replication forks. Here a deoxyribonucleotide (deoxynucleotide triphosphate, or dNTP) polymerizes to form new complementary strands (Figure 32-12). DNA replication occurs bidirectionally from the two replication origin sites. Each DNA strand in the replication fork serves as a template for the formation of a daughter or complementary strand through the activity of DNA polymerase.⁷ The DNA polymerase substrate is the free hydroxyl group located on the 3′ carbon of a deoxyribonucleotide. DNA polymerase recognizes the group and catalyzes the joining of the complementary deoxyribonucleotide. DNA is read 3′ to 5′ by DNA polymerase, and the complementary strand is synthesized 5′ to 3′.

A primer provides the free 3′ hydroxyl group required for DNA polymerase activity. Primers are short nucleotide polymers complementary to the template. The hybridization of the primer to the template requires the enzyme primase. At the replication origin, primase joins a primer to the 3′ end of the 5′-to-3′ (top) template strand (Figure 32-13). Then DNA polymerase recognizes the free hydroxyl group on the 3′ carbon of the last nucleotide in the primer and catalyzes the formation of phosphodiester bonds between the correct complementary nucleotide triphosphate and the primer, releasing the β- and γ-phosphate groups. DNA polymerase continues adding deoxyribonucleotides along the replication fork, going to the left of the replication origin, producing the complementary strand called the leading strand.

The second template strand, called the lagging strand, is also read in the 3′-to-5′ direction. To form a complementary strand, a primer hybridizes to the exposed 3′ end of the replication fork. To proceed in the 5′-to-3′ direction, nucleotides are added in fragments toward the origin of replication. As the left replication fork extends to open more of the template strands for replication, additional primers are hybridized, and DNA polymerase uses the primers to initiate the formation of the complementary strand, continuing until it meets a previously hybridized primer.

DNA polymerase not only joins nucleotides, it also degrades the RNA primers and fills in the correct complementary deoxyribonucleotides. Because the replication of the lagging strand produces many small fragments, it is called discontinuous replication, and the fragments are called Okazaki fragments. Finally, the enzyme ligase joins the discontinuous fragments. The replication fork to the right (downstream) is replicated in the same fashion, although the lagging strand is now formed complementary to the top (5′-to-3′) strand, and the leading strand is formed from the 3′-to-5′ strand; the opposite of the situation described occurs for the left replication fork (Figure 32-14).

The cell cycle is highly regulated. At certain critical points within the cycle, decisions are made to continue or begin cell death via apoptosis. This decision may depend on the state of the DNA replicated (Figure 32-15). Normally, the cell detects errors made during replication and either corrects them or begins apoptosis. This prevents the persistence of daughter cells with genetic errors. If the sensing molecules fail, cell division may continue. Debilitating mutations that mediate cell cycle control may result in tumor formation.

One protein responsible for signaling damaged DNA is p53, a tumor suppressor protein. Damaged cells with increased p53 arrest cell division at G₁, which allows time for DNA repair (Figure 32-16). Cells with mutant p53 are unable to arrest cells in G₁; they continue the process of cell division with damaged DNA.^8–10 If the cell can repair the DNA damage, the cell cycle continues. If the cell damage is too severe, the cell undergoes apoptosis. Hematologic malignancies, such as 21% of chronic myelogenous leukemias (CML),^11–13 23% of chronic lymphocytic leukemias (CLL),^14–16 and 17% of acute lymphoblastic leukemias (ALL),^17–19 are associated with a p53 mutation or deletion (see Box 32-3). In summary, DNA synthesis and accurate cell cycle control demand that the integrity of the nucleotide sequence be maintained during DNA replication.

Isolating RNA from Clinical Specimens

RNA isolation poses greater technical challenges than DNA isolation. Ubiquitous ribonucleases (RNases) degrade RNA. These enzymes are the body’s primary defense against pathogens and are found on mammalian epidermal surfaces; therefore, they contaminate all laboratory surfaces.²² Clinical laboratories that isolate RNA must be RNase free, which necessitates costly precautions and decontamination steps.²³

The isolated RNA includes mRNA, rRNA and tRNA, all of which participate in protein synthesis. Depending on cell type, mRNA may comprise only 3% to 5% of the total cellular RNA; therefore, a large specimen may be needed to obtain adequate mRNA. The mRNA does not represent all the information stored in the DNA, only those genes being expressed. Consequently, mRNA provides quantitative information on the genes being expressed in a cell at the time the specimen is collected.

The steps of RNA isolation are (1) RNA release by cell lysis, (2) RNase inhibition with strong chemical agents such as urea or guanidine isothiocyanate, (3) protein and DNA removal, and (4) RNA precipitation. In step 3, extraction is performed using phenol at a pH of 4, chloroform, and isoamyl alcohol. These separate the DNA and protein into the organic phase, while the RNA remains in the aqueous phase. RNA resists acidic pH, whereas DNA is readily depurinated, because acid cleaves the bond between the purine base and the deoxyribose sugar. Therefore, acidic phenol preferentially isolates and preserves RNA while the genomic DNA (all the DNA) is partitioned along with contaminating proteins, lipids, and carbohydrates. As with DNA, precipitating the RNA from the aqueous phase requires the addition of salt to neutralize the charge of the phosphodiester backbone and ethanol to make the nucleic acid insoluble.24,25

Reverse Transcription Polymerase Chain Reaction for Amplifying RNA

Some hematology analyses require mRNA. Genetically altered mRNA sequences translate to an altered protein. For instance, the Philadelphia chromosome (Ph′), carrying the mutation t(9;22)(q34;q11.2), is present in 95% of CML cases plus 20% of adult ALL and 5% of pediatric ALL cases, and in rare instances in acute myeloid leukemia.33,34 Ph′ results from a reciprocal translocation of the ABL (Ableson) gene on chromosome 9 to the breakpoint cluster region (BCR) of chromosome 22, producing a BCR/ABL hybrid (Figure 32-19).^35–37 Transcription of BCR/ABL produces a chimeric mRNA made up of fragments from both the BCR and ABL genes. Translation generates a fusion protein, tyrosine kinase, that alters normal cell cycle control, which results in unrestrained cell proliferation.³⁸ RT-PCR is performed to detect the chimeric mRNA, thus the mRNA template is preferred to DNA. Although the mutation is present at the DNA level, the position at which the two chromosome sections join is variable, whereas the chimeric mRNA is always the same. Also, the DNA includes untranslated introns, which make the chimera too long to replicate. Physiologic mRNA excision and splicing yields a much shorter target that is more easily amplified.

In RT-PCR, the reverse transcriptase enzyme produces complementary DNA (cDNA) from mRNA present in a total RNA sample extracted from patient blood cells (Figure 32-20). PCR subsequently amplifies the cDNA. The RT-PCR master mix includes an oligo(dT), random, or specific primer; reverse transcriptase; deoxyribonucleotides; primers; the mRNA template; and Taq polymerase.

The first step employs reverse transcriptase and a specialized primer to produce an RNA-cDNA hybrid. The primer, called oligo(dT), is a series of thymine nucleotides. Most mRNAs possess a string of adenine nucleotides on the 3′ end called the polyA tail. The oligo(dT) primer anneals to the polyA tail of mRNA. Reverse transcriptase recognizes the hydroxyl group on the last nucleotide of the primer and reads the mRNA template strand, then adds the correct complementary deoxyribonucleotide. Reverse transcriptase continues up the mRNA template strand, joining the complementary deoxyribonucleotides to the growing cDNA strand to form the mRNA-cDNA hybrid. Subsequently, heat denaturation breaks the hydrogen bonds between the mRNA-cDNA hybrid, separating the two strands. The cDNA strand then acts as a template for replication by DNA polymerase. In the next step, the single-stranded cDNA is amplified as in DNA-based PCR using primers specific for a target sequence in the BCR gene and ABL gene. DNA polymerase extends the primers, forming a double-stranded cDNA of the target chimeric gene. The cycling continues, resulting in millions of copies of the BCR/ABL sequence.39,40

Gel Electrophoresis

Nucleic acid phosphate groups confer a net negative charge. Consequently, in electrophoresis, the rate at which DNA fragments (amplicons) pass through gels is proportional to their mass only and, unlike proteins, not their relative charge. DNA fragment mass is a function of the length in base pairs (bp) or kilobase pairs (kb, 1000 × bp). Fragments are sieved through an agarose or polyacrylamide gel matrix by passing a current through the gel as it is bathed in a buffered conducting salt solution. Electrophoresis gel pore diameter is a function of gel concentration. The pores of an agarose gel are larger than the pores of a polyacrylamide gel. When larger fragments (500 bp to 50 kb) are to be separated, an agarose gel is most effective. For smaller DNA fragments (5 to 1000 bp), a polyacrylamide gel is used.⁴¹

DNA fragments (amplicons), controls, and a mass marker or ladder are pipetted into the sample wells near the anode. Electrical current moves the negatively charged fragments toward the positive electrode. Smaller fragments move faster (farther) than larger fragments. The ladder, composed of fragments of known masses, measured in base pairs or kilobase pairs, runs alongside the sample and control lanes. Amplicon masses of the sample and control are determined by comparing their bands with the bands of the ladder (Figure 32-21).

EtBr, SYBR green, or autoradiography may be employed to detect the controls, size marker, and patient DNA fragments. EtBr, a hydrophobic molecule about the same mass as a purine or pyrimidine base, becomes intercalated between the bases of a DNA double helix or between the bases of a nucleic acid fragment, rendering the nucleic acid fluorescent when exposed to UV light. EtBr may be pipetted directly into the gel as it is prepared or diluted in a buffer and applied after electrophoresis. UV light causes EtBr to fluoresce orange, illuminating the nucleic acid bands. EtBr is a mutagen, so many laboratories substitute the safer SYBR green. SYBR green binds to the minor groove of nucleic acid helices and fluoresces.

For autoradiography, deoxyribonucleotides that are incorporated during the PCR elongation step, usually the adenine nucleotide, are conjugated to a radioactive α-phosphate group ([α³²P]dATP). After electrophoresis, x-ray film is placed over the dried gel. The radioactivity in the amplified DNA fragments exposes the film, producing a banding pattern that is interpreted by the laboratory scientist (Figure 32-22).⁴²

An enzyme-based approach may be used in place of autoradiography, thereby eliminating the health, storage, and disposal concerns associated with radioisotopes. Enzyme-conjugated nucleotides are incorporated during extension; alternatively, probes may be hybridized with enzyme-conjugated nucleotides. The enzymes most commonly used are horseradish peroxidase (HRP) and alkaline phosphatase (AP), which cleave luminol or other synthetic chemiluminescent substrates to release visible light. Light generation is confined to those bands that contain HRP- or AP-conjugated DNAs.

Gel electrophoresis is appropriate when the goal is to determine qualitative target presence or absence. Typical target mutations in hematology are four that raise thrombosis risk: the coagulation factor V Leiden mutation, prothrombin G20210A, MTHFR C677T, and MTHFR A1298T. Three others are mutations that affect the metabolism of the antithrombotic drug warfarin. These three, CYP2C9*2, CYP2C9*3, and VKORC1, are among the first targets of pharmacogenetic testing. Patients positive for the presence of any one or more of these mutations are sensitive to warfarin and require smaller than the usual dosages. A normal warfarin dosage risks causing hemorrhage in warfarin-sensitive patients. Several of these example mutations may also be identified using restriction endonuclease methods.

Restriction Endonuclease Methods

One method to determine whether an amplified target DNA fragment contains a mutation of interest uses enzymes called restriction endonucleases (also known as restriction enzymes). These enzymes are produced naturally in bacteria and are so named because they restrict foreign (phage) DNA from entering and destroying the bacterium. Each restriction enzyme recognizes a specific nucleotide sequence and cuts both strands of the target DNA at the sequence, producing restriction fragments. Recognition sequences can be 4 to 15 nucleotides long. There are hundreds of commercially available restriction endonucleases, which allows recognition of many sequences. The number of restriction fragments produced depends on the number of restriction sites present in the amplified target.43,44 Enzyme action at one restriction site produces two restriction fragments, action at two restriction sites produce three restriction fragments, and so on (Figure 32-23). A restriction enzyme detects even a single base substitution, because the mutation alters its sequence and prevents digestion at the site.

A restriction fragment length polymorphism (RFLP) is a mutation or polymorphism-induced change in the position of the restriction fragment. The factor V Leiden mutation is an excellent example of RFLP. Individuals possessing this mutation have an increased risk of venous thrombosis. The factor V Leiden mutation results from the replacement of guanine with adenine at position 1691 (G1691A) of the coagulation factor V gene.45,46 The mutation alters a site normally detected and cut by the restriction enzyme Mnl1. The wild-type (normal) factor V amplicon is 223 bp long with two Mnl1-specific sites. After PCR and incubation with Mnl1, the wild-type amplicon is cut to three restriction fragments, separable using polyacrylamide gel (Figure 32-24). The fragments are 37, 82, and 104 bp long. The mutant gene generates only two fragments with lengths of 82 and 141 bp. A sample from an individual homozygous for the wild-type gene generates the three anticipate fragments; 37, 82, and 104 bp. A sample from an individual homozygous for the factor V Leiden mutation possesses two copies of the mutated factor V gene and generates only two bands, 82 and 141 bp. A sample from a heterozygous individual possesses one normal and one mutated factor V gene and produces four bands of lengths 37, 82, 104, and 141 bp (see Figure 32-1).

Nucleic Acid Hybridization and Southern Blotting

Another detection method employs a nucleic acid probe designed to hybridize (base-pair) to a selected sequence. Nucleic acid hybridization may be combined with restriction enzyme digestion to make identifications on an electrophoresis gel.47,48

Dr. Edwin Southern developed the classic Southern blot hybridization, which was originally performed using isolated DNA without amplification, but is now performed using PCR-amplified DNA. The Southern blot procedure, which takes 3 days to perform, is now confined to research applications, but pointed the way to RFLP and probe hybridization technology. Restriction endonuclease EcoRI cuts DNA at many sites (Figure 32-25). Gel electrophoresis separates the fragments. The sample is then acid-depurinated to “nick” the fragments. Sodium hydroxide next denatures the DNA, producing single-stranded DNA without changing its nucleotide sequence or two-dimensional position on the gel. The single-stranded DNA is transferred to a nitrocellulose filter by electric current or capillary action, so that the nitrocellulose filter reflects the banding pattern. The DNA is permanently affixed to the nitrocellulose filter by baking and UV cross-linking.^49,50

In the classic Southern blot procedure, detection of the band containing the sequence of interest requires a radioactive or enzyme (horseradish peroxidase or alkaline phosphatase)–conjugated, single-stranded probe complementary to the target sequence. The probe hybridizes to the target DNA, leftover unhybridized probe is washed off, and the hybridized bands are visualized by autoradiography. Southern blot analysis may also be performed using a fluorescently labeled probe instead of a radioactive one.51,52

A time-honored application of the Southern blot technique is detection of B-cell immunoglobulin heavy chain gene or T-cell receptor (TCR) gene rearrangement. Each immunoglobulin heavy chain or TCR gene possesses a unique sequence, the result of somatic rearrangement. Rearrangement joins distinct gene segments from pooled clusters with unique sequences. Thus, each B cell and its progeny produce a specific antibody, and each T cell and its progeny possess a specific cell receptor.53–55 Lymphoproliferative disorders arise from a malignant transformation of a B or T cell. The malignant cells, no longer under the control of cell division regulatory proteins, divide uncontrollably, forming a malignant clone. All the cells within the clone contain the same gene rearrangement; thus the cells are monoclonal. Gene rearrangement analysis detects the monoclonal population and determines whether it is a B-cell or T-cell lymphoproliferative disorder.

The appropriate specimens for gene rearrangement analysis are bone marrow, blood, or tissue from the tumor site. After DNA extraction, restriction enzymes EcoRI, BamHI, and HindIII cut the DNA into restriction fragments. Agarose gel electrophoresis separates restriction fragments. The bands are then depurinated, denatured, and transferred to a nitrocellulose filter. Probes hybridize the nucleotide sequences in either B- or T-cell genes, and autoradiography visualizes the banding pattern. The presence of a distinct band represents a monoclonal cell population. A polyclonal population, consisting of cells with different gene rearrangements, appears as a smear with no distinct banding pattern.56–58 Gene rearrangement analysis provides physicians with an important tool to diagnose and monitor the progress of lymphoproliferative disorders.

Hybridization Labels

Labels are visualizing molecules conjugated to, or incorporated into, probes or primers. Radioactivity is incorporated by introducing isotope-labeled nucleotides during probe synthesis. Radioactivity is detected by applying the sample to photographic film, which is an example of the use of a direct label (Figure 32-26). Direct labeling means that the molecule can be visualized without the addition of a reporter system, whereas indirect labels require additional molecules and reactions to be visualized. For health and disposal reasons, the use of radioactive isotopes is restricted in the clinical setting, but alternative labeling techniques and detection kits are commercially available. For instance, fluorogenic acridinium ester molecules may be incorporated as a direct label. Sodium tetraborate solution, containing 1% Triton X-100, degrades acridinium ester on unhybridized probes, whereas base pairing of the probe to its target protects the ester.⁵⁹ The bound ester is detected by a brief emission of light after addition of hydrogen peroxide.

Biotin and digoxigenin are two commonly used indirect labels. Biotin-labeled probes bind avidin. Avidin is in turn linked to a fluorescent label or to a detectable enzyme (see Figure 32-26). Digoxigenin-labeled probes are bound by an antidigoxigenin antibody conjugated to a fluorescent molecule or enzyme. Alkaline phosphatase or horseradish peroxidase are the enzymes most commonly used in indirect labeling systems. Their activity cleaves substrates to produce visible pigments or chemiluminesence.^60–62

DNA Sequencing

The ability to read the sequence of the nucleic acid has been just as important as PCR in the development of molecular biology.⁶³ A combination of these two important techniques (cycle sequencing) has made DNA sequencing more efficient and its analysis less subjective. In cycle sequencing, the order of the nucleotide bases is determined after amplification.⁶⁴ Cycle sequencing is applied in molecular testing to assess amplified sequences for insertions, deletions, or mutations, such as coagulation factor V Leiden mutation or β-globin mutation.

Cycle sequencing is based on dideoxynucleotide terminator sequencing.⁶⁵ The addition of nucleotides to a growing polymer requires a 3′ hydroxyl group on the last added nucleotide and a triphosphate group on the 5′ end of the next nucleotide to be added. If a nucleotide lacks the 3′ hydroxyl group, it can be incorporated but cannot be added to, so the fragment terminates at the “defective” base (Figure 32-27). If small concentrations of dideoxyadenosine triphosphate, dideoxycytosine triphosphate, dideoxyguanine triphosphate, and dideoxythymine triphosphate each are included in the PCR master mix, over a number of cycles a series of fragments that terminate at each successive base is produced. This is called a nested series of fragments. The dideoxynucleotides are fluorescently labeled to produce a different color for each base, and the fragments are subjected to capillary electrophoresis. The fragments with their labels pass through the beam of a detecting laser one by one, with their order based on their length, which allows their sequence to be read. Because DNA is double-stranded, two PCR primers would produce two series of nested fragments and the detector would read two bases at each position. To avoid this, the PCR reaction is done with one primer, which produces single-sided PCR.

With the sickle cell mutation, for example, sequencing of either, but not both, strands will show whether the mutation (adenine to thymine) is present. Each cell has two copies (alleles) of somatic genes; therefore, sequencing will produce a nested series of fragments from each allele. If the patient is homozygotic, the two nested series of fragments will be identical, whether wild-type or mutant. If the patient is heterozygotic, both wild-type and mutant fragments will be produced in the single-sided PCR, generating two nested series of fragments. In analysis of this sequence, both adenine and thymine signals will be present at the position of the mutation, but because only half the templates contain each sequence, the signals of the adenine and thymine will be half as strong. Sequencing is a reliable method for detection of mutations or single nucleotide polymorphisms in DNA, but it is expensive and requires significant instrumentation, so it is not often used in the clinical setting.

Minimal Residual Disease in Leukemia

Real-time quantitative PCR provides the opportunity to measure minimal residual disease in leukemia, a key indicator of treatment efficacy, clinical remission, and prognosis.71,72 Currently, chemotherapy, radiation therapy, and peripheral blood stem cell transplantation reduce leukemic cells to levels undetectable first by visual bone marrow smear or peripheral blood film review and later by flow cytometry assay.⁷³ This is called minimal residual disease, but despite the seeming disappearance of leukemic cells as recorded by these phenotypic assays, 10¹⁰ malignant cells may continue to reside undetected in blood, marrow, or lymphatic tissue.⁷⁴ Real-time quantitative PCR identifies residual cells and helps guide the types and intensity of therapy with the goal of “molecular” remission. Subsequent to remission, periodic real-time quantitative PCR assays may be used to detect early relapse and drug resistance, enabling the hematologist to initiate follow-up therapy.⁷⁵

Real-time quantitative PCR may detect a single malignant cell within a population of a million cells, providing unparalleled sensitivity. Manufacturers compete to develop series of FRET-labeled probes that are comprehensive, speedy, and specific for various hematologic diseases.⁷⁶ Current applications include detection of BCR/ABL (Ph′) in CML and some acute leukemias (see Figure 31-19); JAK2 (“just another kinase” or Janus kinase) in the myeloproliferative neoplasms polycythemia vera and essential thrombocythemia; the t(15;17)(q22;q21) mutation in acute promyelocytic leukemia; and gene rearrangement in lymphatic neoplasms.⁷⁷

Real-time quantitative PCR may also exploit gene rearrangement in lymphomas or lymphocytic leukemias to test for minimal residual disease. Because the rearranged base sequence varies depending on the individual patient’s malignant clone, the patient’s blood must first be collected and the tumor cell DNA cloned and sequenced.⁷⁸ Thereafter, primers and probes may be designed (and labeled with SYBR green or FRET probes) to replicate and detect the patient’s sequence.⁷⁹ This process is called allele-specific oligonucleotide PCR.

Although no purpose is served by quantitative measurement, the specificity and speed of real-time PCR mean that it is effective for the qualitative detection of germ-cell mutations such as the factor V Leiden mutation, prothrombin G20210A, MTHFR C677T, MTHFR A1298T CYP2C9*2, CYP2C9*3, and VKORC1.

Infectious Disease Load

Real-time quantitative PCR can detect and quantitate a number of blood-borne viruses: hepatitis B and C viruses, human papillomavirus, CMV, Epstein-Barr virus, and HIV.⁸⁰ Human bacterial pathogens such as β-hemolytic streptococcus from throat swabs, anaerobes from wound swabs, and bacteria from urine or other body fluids can be detected within hours of collection. Antibacterial therapy can be initiated based upon the speedy results of molecular susceptibility testing. Real-time quantitative PCR is the reference method for detection and quantification of methicillin-resistant Staphylococcus aureus (MRSA), vancomycin-resistant enterococcus, and opportunistic Clostridium difficile. Molecular diagnostic techniques are effecting in identifying and monitoring malarial and other blood-borne parasites. The challenge to primer and probe developers is to select sequences that are specific enough to avoid false positives caused by nonpathogenic strains, sensitive enough to positively identify infectious strains, and flexible enough to remain effective as pathogenic microorganisms mutate and evolve.

Clinical relevance is important when assessing infectious disease using molecular techniques. Such methods allow millions of copies to be generated from a single DNA sequence from a microorganism or virus. Theoretically, the presence of a single organism can lead to a positive test result, but a single organism may not be clinically relevant when monitoring viral loads. Standard curves of template number are crucial to data interpretation. Also, because DNA survives the organism, a positive result on a test for a given sequence does not guarantee that the organism was viable at the time of sampling.