Rodak's Hematology: Clinical Principles and Applications, 5th Ed.

CHAPTER 31. Molecular diagnostics in hematopathology

Cynthia L. Jackson, Shashi Mehta*


Structure and Function of DNA

The Central Dogma: DNA to RNA to Protein

DNA at the Molecular Level

Transcription and Translation

DNA Replication and the Cell Cycle

Molecular Diagnostic Testing Overview

Nucleic Acid Isolation

Isolating DNA from Clinical Specimens

Isolating RNA from Clinical Specimens

Amplification of Nucleic Acids by Polymerase Chain Reaction

Polymerase Chain Reaction for Amplifying DNA

Reverse Transcription Polymerase Chain Reaction for Amplifying RNA

Detection of Amplified DNA

Gel Electrophoresis

Restriction Endonuclease Methods

Nucleic Acid Hybridization and Southern Blotting

Cleavage-Based Signal Amplification

DNA Sequencing

Real-Time Polymerase Chain Reaction

Qualitative Real-Time Polymerase Chain Reaction

Quantitative Real-Time Polymerase Chain Reaction

Minimal Residual Disease in Leukemia

Mutation Enrichment Strategies

Chromosome Microarrays

Pathogen Detection and Infectious Disease Load

Current Developments


After completion of this chapter, the reader will be able to:

1. Describe the structure of DNA, including the composition of a nucleotide, the double helix, and the antiparallel complementary strand orientation.

2. Predict the nucleotide sequence of a complementary strand of DNA or RNA given the nucleotide sequence of a DNA template.

3. Explain the relationship between DNA structure and protein structure.

4. Discuss the process of DNA replication, including replication origin, replication fork, primase, primer, DNA polymerase, Okazaki fragments, leading strand, and lagging strand.

5. Determine the appropriate patient specimen required for DNA isolation to identify an inherited or somatic mutation.

6. Discuss common methods of DNA and RNA isolation.

7. Explain the principle of polymerase chain reaction (PCR), reverse transcriptase PCR, and nucleic acid hybridization.

8. Compare and contrast the methods for detecting target DNA including: gel electrophoresis using intercalating dyes or capillary gel electrophoresis; cleavage-based signal amplification; restriction fragment length polymorphism; and probe hybridization techniques such as Southern blotting.

9. Describe the principle of qualitative real-time PCR and contrast it with endpoint PCR.

10. Discuss the use of quantitative real-time PCR for monitoring minimal residual disease.

11. Explain traditional Sanger DNA sequencing and contrast it with pyrosequencing, and next-generation sequencing (massively parallel sequencing).

12. Describe the use of microarray-based comparative genomic hybridization (aCGH) and single nucleotide polymorphism array (SNP-A) karyotyping for the detection of chromosome copy number alterations.


After studying the material in this chapter, the reader should be able to respond to the following case study:

A 42-year-old man came to the emergency room complaining of pain behind his right knee. He had observed swelling below the knee for the previous 2 days. The patient was in no apparent distress and was experiencing no chest pain, shortness of breath, dyspnea, or hemoptysis. The patient reported no history of trauma except for a right femur break 20 years before. He reported that he was taking no medications and was in good general health. Five years previously, he experienced an episode of deep vein thrombosis (DVT) in his right lower leg and was treated with intravenous heparin followed by oral warfarin (Coumadin) for 3 months. Subsequent to treatment, he experienced occasional pain behind both knees, which he treated with aspirin. He noted that his mother had been diagnosed with carpal tunnel syndrome and had developed DVT, for which she has been taking oral warfarin for 15 years. The patient’s job requires frequent long airplane flights. He flies first class and walks around occasionally during the long flights. His leg pain began 1 week after a flight to Europe.

On physical examination, the patient had no evidence of rash or oral ulcers. No petechiae or purpura were noted. He had mild pretibial pitting edema. His right leg measured 36.5 cm at 25 cm distal to the superior aspect of the patella, whereas his left leg measured 33.5 cm in the same location. CBC findings were unremarkable, and both the prothrombin time and activated partial prothrombin time were within the reference intervals. Doppler ultrasonography revealed complete occlusion of the distal superficial femoral vein, anterior tibial vein, and popliteal vein. The diagnosis was DVT without pulmonary emboli. The patient was hospitalized, and a heparin drip was started. The hematologist ordered a factor V (F5) Leiden mutation analysis on blood drawn in an EDTA tube. Figure 31-1 illustrates the results of the mutation analysis.

1. What type of specimen is appropriate when analyzing DNA for a hereditary mutation?

2. Examine the gel electrophoresis result (Figure 31-1). Are the correct controls present?

3. What band sizes (in base pairs or bp) appear in the patient’s sample?

4. What band sizes are expected for an individual who is homozygous for the factor V (F5) Leiden mutation, heterozygous for the mutation, and free of the mutation?

5. Does this patient have the factor V (F5) Leiden gene mutation?

**This case was provided by George A. Fritsma, MS, MLS, manager, The Fritsma Factor: Your Interactive Hemostasis Resource,, sponsored by Precision BioLogic, Inc., Cambridge, Nova Scotia.


FIGURE 31-1 Results of the factor V (F5) Leiden mutation test on the patient in the case study. Lane A, molecular size marker; B, positive control (homozygous for factor V Leiden mutation); C, patient’s sample; D, negative control (no mutation); E, no-DNA control. The expected banding pattern on an agarose gel for the factor V (F5) Leiden mutation test is as follows: homozygous for the mutation, 141 and 82 bp bands; heterozygous for the mutation, 141, 104, and 82 bp bands; no mutation (normal or wild-type), 104 and 82 bp bands. A band at 37 bp is barely visible and is difficult to detect on an agarose gel. However, this band is not essential for interpreting the results.

Molecular biology techniques enhance the diagnostic team’s ability to predict or identify an increasing number of diseases in the clinical laboratory. Molecular techniques also enable clinicians to monitor disease progression during treatment, make accurate prognoses, and predict the response to therapeutics. The short interval required to perform molecular diagnostic tests and analyze their results is an additional positive aspect of this type of testing, resulting in more efficient patient management, especially in cases of infection. The five main areas of hematopathologic molecular testing include detection of mutations, gene rearrangements, and chromosomal abnormalities for diagnosis and prognosis of hematologic malignancies (); detection and quantification of minimal residual disease to monitor treatment of hematologic malignancies; detection of mutations in inherited hematologic disorders (Box 31-1Box 31-2); pharmacogenetic testing to detect genetic variation affecting certain drug therapies (Box 31-3); and identification of hematologically important infectious diseases (Box 31-4).

BOX 31-1

Major Hematologic Malignancies in Which Molecular Methods Are Performed for Diagnosis and Monitoring Minimal Residual Disease

For Diagnosis:

Acute leukemias



Myeloproliferative neoplasms

Chronic myelogenous leukemia

Polycythemia vera

Essential thrombocythemia

Primary myelofibrosis

Myelodysplastic syndromes

Mature lymphoid neoplasms

Chronic lymphocytic leukemia


For Monitoring Minimal Residual Disease:

Acute leukemias

Quantification of fusion mRNA transcripts due to translocations

Quantification of specific B and T cell receptor rearrangements

Chronic myelogenous leukemia

Quantification of fusion mRNA transcripts due to translocation

BOX 31-2

Inherited Hematologic Disorders Detected by Molecular Methods

Erythrocyte disorders


Membrane abnormalities

Enzyme deficiencies

Erythropoietic porphyrias

Leukocyte disorders

Quantitative disorders

Functional disorders

Storage disorders

Platelet disorders

Quantitative disorders

Functional disorders

Bone marrow failure syndromes



BOX 31-3

Pharmacogenetic Testing for Genetic Variation Affecting Therapy

Warfarin sensitivity

Cytochrome P450 2C9 variants, CYP2C9*2, CYP2C9*3

VKORC1 variants

Clopidogrel sensitivity

Cytochrome P450 2C19 variants, CYP2C19*17, others

Thiopurine sensitivity

Thiopurine S-methyltransferase, TPMT*2, TPMT*3C, TPMT*3A

Imatinib resistance

ABL1 mutation analysis

BOX 31-4

Hematologically Important Pathogens Detected by Molecular Methods

Parasitic pathogens






Fungal pathogens

Bacterial pathogens

Viral pathogens

Parvovirus B19


Epstein-Barr virus

Human immunodeficiency virus types 1 and 2

Human T-cell lymphotropic virus type 1

Modified from Paessler M, Bagg A: Use of molecular techniques in the analysis of hematologic diseases. In Hoffman R, Benz EJ Jr, Shattil SJ, et al, editors: Hematology: basic principles and practice, ed 4, Philadelphia, 2005, Churchill Livingstone, pp. 2713-2726.

Structure and function of DNA

The central dogma: DNA to RNA to protein

Much of the stored information needed to carry out cell processes resides in deoxyribonucleic acid (DNA); therefore, proper cellular storage, maintenance, and replication of DNA are necessary to ensure homeostasis. Because molecular testing takes advantage of DNA structure and replication, a review of molecular biology is helpful.

The central dogma in genetics is that information stored in the DNA is replicated to daughter DNA, transcribed to messenger ribonucleic acid (mRNA), and translated into a functional protein (Figure 31-2). This process is essential to carry out cellular functions while preserving a record of the stored information. In eukaryotes, the initial DNA sequence is composed of translated exons separated by untranslated introns. The introns are enzymatically excised following transcription from DNA to RNA, and the mature mRNA sequence is then translated. Translation is an enzymatic process wherein mRNA three-nucleotide base sequences called codons drive the addition of individual amino acids to the growing peptide. The mature protein then carries out its cellular function, which may be structural or may involve recognition, regulation, or enzymatic activity.


FIGURE 31-2 RNA polymerase binds to a sequence of DNA called the promoter region, which causes the DNA strands to separate. Using one of the DNA strands as a template, RNA polymerase moves along and simultaneously reads the DNA strand, forming the primary messenger RNA (mRNA) transcript by joining the complementary ribonucleotides. The primary mRNA transcript consists of sequences called exons that provide coding information and introns that are excised from the mature mRNA. The spliced mRNA then leaves the nucleus and enters the cytoplasm of the cell where the ribosomes translate the mRNA into protein.

The structural units that carry DNA’s message are called genes. The human β-globin gene, part of the hemoglobin molecule, provides a good example of replication and transcription, because it was one of the first sequenced and demonstrates the result of aberrant sequence maintenance. A normal (or wild-type) β-globin gene contains a sequence of bases that code for a β-globin peptide of 146 amino acids (Chapter 10). One inherited mutation changes a single DNA base. This is called a point mutation. The mutation occurs in the portion of the sequence that codes for the sixth amino acid of β-globin. The mutation substitutes the amino acid valine for glutamic acid in the growing peptide. Valine modifies the overall charge, producing a protein that polymerizes in a low-oxygen environment. This leads to sickled erythrocytes, circulatory ischemia, and poor oxygen exchange between blood and tissues.12 A mutation in one of the two copies (alleles) of this gene inherited from the parents results in a heterozygous condition, or sickle cell trait. In a heterozygote,the symptoms of the disease are often unseen or are present only during times of physical stress. If both alleles are mutated, there is overt homozygous sickle cell disease, and the symptoms are severe.

Every active gene is translated. Human somatic cells contain 20,000 to 25,000 genes in 2 meters of DNA.34 Significant packing (Figure 30-3) takes place to reduce the volume of the nucleic acid to the size of chromosomes.

DNA at the molecular level

DNA is a duplex molecule composed of two complementary hydrogen-bonded nucleotide strands (Figure 31-3). 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. 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 (Figure 31-4A). 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.


FIGURE 31-3 DNA is a double-stranded helical macromolecule consisting of nucleotide subunits joined in sequence by deoxyribose molecules (pentagons) and phosphate radicals (circles). The bases thymine (T), adenine (A), cytosine (C), and guanine (G) are illustrated in their standard pairs: thymine to adenine, cytosine to guanine.


FIGURE 31-4 A, The pentose sugar, deoxyribose, a phosphate group, and a nitrogenous base compose a DNA nucleotide. The carbons of the deoxyribose molecule are numbered 1′ through 5′. The hydroxyl group on the 2′ carbon of ribose is replaced by a hydrogen molecule, making the structure a deoxyribose. B, A nucleotide results from the formation of a glycosidic bond between the nitrogenous base and the hydroxyl group on the 1′ carbon of deoxyribose and a phosphodiester bond between the phosphate group and the hydroxyl group on the 5′ carbon of deoxyribose. C, A nucleotide illustrating the glycosidic and phosphodiester bonds.

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 the sugar at the 5′ carbon by a phosphodiester bond (Figure 31-4B, C). 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 with the α-phosphate linked to the sugar (Figure 31-5).


FIGURE 31-5 The enzyme DNA polymerase catalyzes the reaction between the hydroxyl group on the 3′ carbon of one nucleotide with the phosphate group bound to the 5′ carbon of the downstream nucleotide. The α-phosphate group is split by the 3′-OH, with release of the β- and γ-phosphates.

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 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 31-5). Polymerization of subsequent nucleotides forms a DNA strand.

DNA consists of two strands that are antiparallel and complementary (Figure 31-6). One strand begins with a phosphate group attached to the 5′ carbon of the first nucleotide and ends with the hydroxyl group on the 3′ carbon of the last nucleotide. 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.


FIGURE 31-6 DNA consists of two antiparallel and complementary strands. One strand begins with a 5′ phosphate group and ends with a 3′ hydroxyl group. This strand is read in the 5′-to-3′ direction. The other strand begins with a 3′ hydroxyl group and ends with a 5′ phosphate group. This strand is shown in the 3′-to-5′ orientation.

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 31-7). 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 31-8). RNA is most often single-stranded but can have significant secondary structure.


FIGURE 31-7 The single-ringed pyrimidines (thymine and cytosine) and the double-ringed purines (adenine and guanine) are the code-carrying nitrogenous bases of DNA.


FIGURE 31-8 A, The purine adenine forms two hydrogen bonds with the pyrimidine thymine. The purine guanine forms three hydrogen bonds with the pyrimidine cytosine. B, The two strands maintain a consistent distance from each other, which allows DNA to twist into a helix.

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 which separates the DNA strands. The enzyme then slides along the 3′ to 5′ template DNA strand, “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 31-2). If the nucleotide sequence of the template DNA strand is 3′-CTAG-5′, the primary RNA transcript is 5′-GAUC-3′, where uracil is substituted for thymine.

Primary mRNA transcripts are composed of introns and exons. Introns are untranslated intervening sequences located between 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 (polyA tail).6 The mature mRNA leaves the nucleus and enters cytoplasm to be translated by the ribosomes.

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 codon is reached. Three termination codons exist that do not code for any amino acid and terminate translation: UAA, UAG, and UGA. 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 (Chapter 6)The cell cycle progresses through a defined sequence (Figure 31-9). 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. Checkpoints occur at the end of G1 before DNA replication in the S phase, and at the end of G2 before mitosis in the M phase. The checkpoints have complex mechanisms to stop the progression of the cell cycle if a problem is detected, at which point the cell will undergo apoptosis. 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.


FIGURE 31-9 The cell cycle consists of interphase and mitosis. Interphase is divided into G1, S, and G2 phases. Cell growth occurs during G1. During the S phase, DNA synthesis or replication occurs. The cell prepares for mitosis during the G2 phase. During mitosis the cell divides, producing two identical daughter cells. The cells may also enter a quiescent phase called G0, where the cell functions but does not divide. There are two critical times in the cell cycle that are checkpoints where the cell will either continue through the cell cycle or undergo apoptosis. The first checkpoint is before S phase and DNA replication, and the second checkpoint is at the end of G2, where the cell will either enter mitosis or undergo apoptosis.

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 the hydrogen bonds, and untwists and separates the DNA strands producing two replication forks. Here a deoxyribonucleotide (deoxynucleotide triphosphate, or dNTP) polymerizes to form new complementary strands (Figure 31-10). 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 grouplocated on the 3′ carbon of a deoxyribonucleotide. DNA polymerase recognizes this group and catalyzes the joining of the complementary deoxyribonucleotide. DNA polymerase reads the DNA template in the 3′-to-5′ direction, and the complementary strand is synthesized in the 5′-to-3′ direction.


FIGURE 31-10 DNA Replication. A, Primases synthesize RNA primers that anneal to the single-stranded template strands. The primers must be oriented in such a way that the hydroxyl group on the 3′ end of the primers is available for deoxyribonucleotide addition by DNA polymerase. B, DNA polymerase extends the primer located on the 5′-to-3′ coding strand, producing the complementary leading strand (blue). On the 3′-to-5′ template strand, DNA polymerase extends the primers, producing Okazaki fragments. The primer ribonucleotides (red) are replaced with deoxynucleotides by DNA polymerase to produce the complementary lagging strand (green). C, Bidirectional DNA replication shown in which the 5′-to-3′ parent strand serves as the template for producing the continuous leading strands on a replication fork to the left of an origin. The 3′-to-5′ parent strand is the template for the lagging strands, which are produced in a discontinuous manner. The continuous and discontinuous strands are reversed on the replication fork to the right.

primer is required to provide the free 3′ hydroxyl group that is necessary for DNA polymerase activity. The enzyme primase synthesizes short RNA polymers complementary to the template that serve as primers to initiate DNA synthesis. At the replication origin, the primer hybridizes to the 3′ end of the 5′-to-3′ (top) template strand (Figure 31-10). 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, but 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 31-10).

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. 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. In summary, DNA synthesis and accurate cell cycle control demand that the integrity of the nucleotide sequence be maintained during DNA replication.

Molecular diagnostic testing overview

DNA or RNA sequences are used to diagnose and monitor solid tumors, acute leukemia, myeloproliferative disorders, myelodysplastic neoplasms, inherited thrombosis risk factors, and viral, parasitic, and bacterial infections. Molecular diagnostic testing exploits the enzymes and processes of DNA replication. Most molecular testing methods use replication—for example, polymerase chain reaction (PCR)—to make millions of amplicons (copies) of a DNA sequence of interest. Further, creation of synthetic DNA requires the use of short sequences used as either primers or probes to locate specific DNA or RNA sequences within vast populations of nucleic acids.

Specific mutations are associated with hematologic disease. These are detected by allele-specific amplification methods, DNA sequencing, or restriction fragment length polymorphism analysis of amplified material. Messenger and ribosomal RNA also may be amplified through a process called reverse transcriptase PCR (RT-PCR). Using mRNA, the existence of mutations that are being actively translated can be detected. Assessment of mRNA shows whether a mutation is expressed in a certain cell type or tissue and can be used to quantitatively determine the level of transcription of a gene. It can also be used to detect and monitor chromosome translocations that produce novel chimeric mRNA transcripts in conditions where the breakpoints are too widely separated to be detected by PCR amplification of DNA.

Most molecular tests use DNA amplification such as PCR, generating multiple amplicons of the target sequence. Amplification is meant to be specific to the sequence of interest in the sample being tested; however, it will amplify any DNA that is present in the reaction. Consequently, it is critical to eliminate contamination of newly isolated target DNA with amplicons from previously amplified samples. Contamination can be avoided by designating separate laboratory locations for each step, having a unidirectional work flow, and employing appropriate controls. Operators routinely employ ultraviolet (UV) light and bleach to induce strand breaks in contaminating DNA on work surfaces and a uracil-N-glycosylase system that destroys previously amplified DNA can also be incorporated into the PCR reactions.

In genetically based hematologic disease, mutations and polymorphisms can occur that do not affect function. Individuals vary in genetic sequences coding for identical proteins. Such single nucleotide polymorphisms are commonly detected but might not be associated with disease. With these caveats in mind, several techniques are presented and an example from hematopathology is given for each. is a summary of molecular methods with hematopathology applications. Box 31-5

BOX 31-5

Molecular Methods with Hematopathology Applications

Nucleic acid isolation



Amplification of nucleic acid

Polymerase chain reaction (PCR)

Reverse transcriptase PCR

Detection of amplified DNA


Restriction endonuclease methods

Nucleic acid hybridization and Southern blotting

Cleavage-based signal amplification

DNA sequencing

Real-time PCR



Minimal residual disease in leukemia

Mutation enrichment strategies

Chromosomal microarrays

Pathogen detection and infectious disease load

Current developments

Mass spectrometry

Digital PCR

Next-generation sequencing

Nucleic acid isolation

Isolating DNA from clinical specimens

Most molecular diagnostic tests begin with the isolation of DNA or RNA from a patient specimen. To test for a mutation in patient DNA, the patient’s DNA is isolated. To test for microorganism DNA, as in an infection, DNA is also isolated from the patient specimen because it will include the organism DNA. The preferred nucleic acid for clinical diagnosis is DNA because it is inherently more stable than RNA and is less labor intensive to isolate.

The molecular laboratory isolates nucleic acid from a wide variety of clinical specimen types. Patient specimens for human DNA isolation may include peripheral blood, bone marrow, tissue biopsy specimens (both fresh and formalin fixed paraffin-embedded), needle aspirates, body fluids, saliva, and cheek swabs. Blood, saliva, or cheek swab specimens are all appropriate for identifying an inherited defect, although blood is the most common specimen type. Every nucleated cell contains a full complement of DNA. If individuals inherit a mutation, it is present in the DNA of all their nucleated cells, both gamete and non-gamete (somatic) cells. Thus the DNA in the nucleus of white blood cells can reveal inherited mutations. In solid tumors, somatic (acquired) mutations are detected by analyzing DNA from the suspect tissue. For identification of infectious disease organisms by molecular techniques, DNA must also be isolated from the affected tissues. Peripheral blood is adequate for infections with viruses such as human immunodeficiency virus (HIV) and cytomegalovirus (CMV) that infect blood cells, whereas cerebrospinal fluid is required for meningeal infections.

Whole blood is preferentially collected in an ethylenediaminetetraacetic acid (EDTA) tube to prevent clotting and to inhibit enzymes that may digest DNA, although other tubes may also be acceptable. The red blood cells (RBCs) are removed by taking advantage of the differential lysis in hypotonic buffer due to differing osmotic fragility between white and red blood cells. Incubation in hypotonic buffer will result in the red blood cells lysing before the white blood cells, thus allowing the WBCs to be removed from the hemoglobin and lysed RBCs by centrifugation. Hemoglobin is a potent inhibitor of PCR and other downstream procedures.8

DNA from tissue suspected of being cancerous can be isolated from formalin-fixed, paraffin-embedded tissue sections mounted on glass microscope slides or whole sections cut directly into a microfuge tube. Tissue is obtained from the entire section or from a portion of the section by microdissection, either by scraping or by laser. The tissue is degraded by an enzyme called proteinase K to break open the cells and release the DNA. The sample is then purified using an automated or manual extraction kit as described below.9 In addition to paraffin-embedded samples, fresh or frozen tissue samples are also appropriate for DNA isolation. Quickly thawing and mincing the frozen tissue prepares the sample for DNA isolation. The minced tissue is mixed with an extraction buffer to release the DNA from the cells, and it is then purified.

There are a number of automated extraction systems as well as manual extraction kits available for DNA extraction. Most of these systems use a solid phase extraction system that takes advantage of the binding of DNA to silica under high salt conditions. Manual kits use columns that can be spun in a microcentrifuge with the eluent collected in microfuge tubes. Cells that have been lysed and protease treated are applied to a column in a high salt buffer. The column is washed to remove impurities and the DNA eluted in a low-ionic-strength buffer and collected in a microfuge tube.10 Automated extractors have reagents packaged in sets and can be programmed to extract and purify the DNA automatically. There are a variety of models to choose from, depending on the number and type of samples. Isolated DNA can be stored at –20°C. If a delay in the molecular testing is necessary, the isolated DNA sample can be stored at −80° C indefinitely.

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.11 Clinical laboratories that isolate RNA must be RNase free, which necessitates costly precautions and decontamination steps.12

The isolated total 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.

RNA may be purified using either liquid or solid phase procedures. The steps of RNA isolation using a liquid phase method are (1) RNA release by cell lysis combined with RNase inhibition by homogenization or incubation in a strongly denaturing solution containing chemical agents such as urea or guanidine isothiocyanate, (2) protein and DNA removal, and (3) RNA precipitation using alcohols. In step 2, extraction is performed using acidic phenol chloroform and guanidine isothiocyanate.

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. 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.1314 Purification of RNA using column-based methods is similar to DNA except that the RNA is suspended in a high-salt buffer that preferentially binds RNA greater than 200 nucleotides to the column to remove smaller RNAs such as tRNA and 5S RNA.

Amplification of nucleic acids by polymerase chain reaction

Polymerase chain reaction for amplifying DNA

Polymerase chain reaction (PCR) is the principal technique in the clinical molecular laboratory. PCR is an enzyme-based method for amplifying a specific target sequence to allow its detection from a small amount of highly complex material.15 Sickle cell anemia results from a single β-globin nucleotide substitution (point mutation) in which an adenine replaces a thymine. Detecting this mutation from among 6 billion nucleotides in the human genome would be like finding a needle in a haystack if only a few cells were assessed. When millions of β-globin copies are produced, however, the mutation is easily detected.16There are two categories of PCR reactions: endpoint PCR and real-time PCR. The amplification for both categories is basically the same. The major difference is in the method of detection of the PCR product. With endpoint or standard PCR, the amplification products must be detected using another technique such as gel electrophoresis, which is described later in the chapter. In real-time PCR, the amplicons are detected during the PCR cycles by using fluorescence detection. This is also described in further detail later in the chapter.

As with natural DNA replication, PCR amplification requires primers that anneal (bind) to complementary nucleotide sequences on either side of the target region. In testing for the sickle cell mutation, for example, selected primers flank (i.e., bind on either side of) the β-globin gene sequence containing the mutation. The total base pair (bp) length of the primer sequences plus the target sequence can vary, but in this example, it is 110 bp for the β-globin gene, a typical sequence length for many mutation sites ().Figure 31-1116 Besides primers, the PCR master mix reagents include a heat-insensitive DNA polymerase—for example, Taq polymerase—isolated from the thermophilic bacterium T hermus aq uaticus and a mixture of the four deoxyribonucleotides—deoxyadenosine triphosphate (dATP), deoxythymidine triphosphate (dTTP), deoxyguanosine triphosphate (dGTP), and deoxycytidine triphosphate (dCTP)—in a magnesium buffer.


FIGURE 31-11 Application of PCR to target ß-globin DNA. PCR amplifies the target DNA, making millions of copies of the target DNA after 30 cycles. Flanking forward and reverse primers (PCO3 and PCO4) are used to amplify the target ß-globin DNA. One primer (PCO3, orange) anneals to the 3′ end of the 3′-to-5′ DNA strand. The other primer (PCO4, green) anneals to the 3′ end of the 5′-to-3′ DNA strand. These primers provide the 3′-OH end for extension in the 5’ to 3’ direction during the PCR reaction and set the boundaries for the size of the amplicon. dsDNA, double-stranded DNA; ssDNA, single-stranded DNA.

The DNA is first denatured at 95° C, which separates the strands; then cooled to the primer annealing (binding) temperature of 40° to 60° C; and then warmed to 72° C to promote specific chain extension, in which nucleotides are added to the primers by DNA polymerase. The annealing temperature is optimized for each set of primers. A thermocycler is used to accurately produce and monitor the rapid temperature changes.

Once the double-stranded DNA is denatured, one primer anneals to the 3′ end of the 5′-to-3′ strand and the other primer to the 3′ end of the complementary 3′-to-5′ strand. Both primers possess a free 3′ hydroxyl group. The DNA polymerase recognizes this hydroxyl group, reads the template, and catalyzes formation of the phosphodiester bond joining the first complementary deoxyribonucleotide to the primer. The polymerase rapidly continues down the template strand at 1000 nucleotides per second, extending the complementary strand in the 5′-to-3′ direction to eventually produce a complete daughter strand that continues to the 5′ end of the template.17 This completes one PCR cycle. In the second cycle, the temperature changes are repeated, and the first-cycle product becomes the template for a daughter strand. After the second cycle, daughter strands are produced that are bounded by the primer sequences at the 5′ and 3′ ends, resulting in a fragment of DNA of the desired length. In 25 to 40 subsequent cycles, this DNA of specific length and sequence, called an amplicon, is reproduced millions of times.1819

Primer annealing accounts for PCR specificity, and primer design is crucial for achieving confidence in the test results regardless of the application. Wherever primers anneal, specifically or nonspecifically, they become starting points for extension.

Commercial kits contain primer sets that have been tested for annealing specificity, but care must be taken to use the optimal annealing temperature. Even if the primer is properly designed, it can anneal to noncomplementary regions if the annealing temperature is too low. Several online primer design programs are available from genome centers and company websites. One such program that can help determine the uniqueness and therefore specificity of the primers is the Basic Local Alignment Sequence Tool (BLAST).20 These programs will also analyze pairs of primers to avoid complementarity between the primers themselves to prevent hybridization to one another, which forms undesirable primer dimers.

Controls are essential for the accurate interpretation of a PCR result. The three controls required for PCR are the negative, positive, and “no-DNA” or no template (NTC) controls. All three are included in each run. In addition, in most applications, a sensitivity control will be included that consists of a low positive sample at the lowest concentration detected. The negative control consists of DNA known to lack the sequence of interest; the positive control contains the target sequence. Comparison of the amplification in the patient sample to results in the negative and positive controls determines whether the target DNA sequence is present in the patient’s DNA. The no-DNA control detects master mix contamination. Amplification in the no template control indicates DNA contamination, which renders the entire test result unreliable.21

Reverse transcription polymerase chain reaction for amplifying RNA

Some hematology molecular tests such as those for translocations, require mRNA as the starting material. Genetically altered mRNA sequences often translate to an altered protein. The classic example is the Philadelphia chromosome (Ph′), carrying the chromosome translocation t(9; 22)(q34; q11.2) (Chapter 33). This translocation is present in 95% of chronic myelogenous leukemia (CML) cases, as well as 20% of adult acute lymphoblastic leukemia (ALL) and 5% of pediatric ALL cases, and in rare instances in acute myeloid leukemia.2223 Ph′ results from a reciprocal translocation of the ABL1 (Ableson) gene on chromosome 9 to the breakpoint cluster region (BCR) of chromosome 22, producing a BCR-ABL1 hybrid or chimeric gene (Figure 31-12A). Transcription of BCR-ABL1 produces a chimeric mRNA made up of exons from both the BCR and ABL1 genes. Translation generates a fusion protein, tyrosine kinase, that alters normal cell cycle control, which results in unrestrained cell proliferation.27 RT-PCR of the chimeric mRNA is the standard method to detect this mutation. Although the mutation is present at the DNA level, the nucleotide position at which the two chromosome sections join is variable, whereas the chimeric mRNA is always the same. The DNA also includes untranslated introns, which make the chimera too long to replicate. The physiologic excision and splicing of mRNA yields a much shorter target that is more easily amplified.



FIGURE 31-12 A, The BCR gene is present on chromosome 22, and the ABL1 gene is located on chromosome 9. The Ph′ chromosome results from the translocation of the ABL1 gene to chromosome 22, which places the ABL1 gene next to the BCRgene and produces a chimeric BCR/ABL1 gene. The transcription of the BCR/ABL1 gene produces a chimeric messenger RNA (mRNA) consisting of a portion of the BCR gene and a portion of the ABL1 gene. B, Reverse transcriptase polymerase chain reaction (RT-PCR) produces complementary DNA (cDNA) from messenger RNA (mRNA). This diagram shows the RT-PCR steps used to produce amplified BCR/ABL1 cDNA. Initially, a gene-specific primer or short random primers anneal to the chimeric BCR/ABL1 mRNA. Reverse transcriptase elongates the primer, producing an mRNA-cDNA hybrid. Heat denaturation breaks the hydrogen bonds, holding the hybrid molecule together, releasing the single-stranded (ss) BCR/ABL1 cDNA. Next, a primer specific for the ABL1 gene is annealed to the cDNA. DNA polymerase elongates the primer, producing the double-stranded (ds) BCR/ABL1 cDNA. The cDNA becomes single stranded by heat denaturation. Then the ABL1 primer as well as a primer specific for the BCR gene anneal to the ss cDNA. DNA polymerase elongates the primers, producing ds BCR/ABL1 cDNA. The cycle is repeated 20 to 40 times, producing millions of copies of the ds BCR/ABL1 cDNA.

In RT-PCR, the reverse transcriptase enzyme produces complementary DNA (cDNA) from mRNA present in a total RNA sample extracted from patient specimens such as blood or bone marrow (Figure 31-12). PCR subsequently amplifies the cDNA.

The first step is to transcribe the RNA into DNA using reverse transcriptase and a primer to produce an RNA-cDNA hybrid. The primer can be oligo(dT), a series of thymine nucleotides complementary to the string of adenine nucleotides on the 3′ end of most mRNAs, called the polyA tail; a set of short random primers that prime the cDNA synthesis more evenly; or a specific primer for the gene of interest. The primer anneals to the complementary sequence of the 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 along 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. The cDNA synthesis can be done separately from the PCR amplification step in a two- step procedure or combined with the PCR in a single reaction. For example, with the BCR-ABL1 translocation, the single-stranded cDNA is amplified as in DNA-based PCR using one primer specific for a target sequence in the BCR gene and a second primer specific for the ABL1 gene. DNA polymerase extends the primers, forming a double-stranded cDNA of the target chimeric gene. Only the cDNA containing the translocation, and therefore both primer binding sites, will be amplified, resulting in millions of copies of the BCR-ABL1sequence.2829

Detection of amplified DNA

Although many molecular tests are now performed using real-time PCR, there are still circumstances where amplicons are produced using endpoint PCR and the product must be detected using downstream techniques. Amplified target DNA may be detected by gel electrophoresis using fluorescent dyes. PCR can also be combined with restriction enzyme digestion of the amplicons followed by gel electrophoresis or cleavage-based signal amplification (Invader) technology for detection (discussed later in the chapter).

Gel electrophoresis

Nucleic acid phosphate groups confer a net negative charge to DNA fragments. Consequently, in electrophoresis, the rate at which DNA fragments (amplicons) migrate 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.30

In slab gel electrophoresis, PCR products (amplicons) of patients and controls, and a mass marker or ladder are pipetted into the sample wells in the gel slab near the negative electrode (cathode). An electrical current moves the negatively charged fragments toward the positive electrode (anode). Smaller fragments move faster and migrate farther than larger fragments. The ladder, composed of fragments of known masses (sizes), measured in base pairs or kilobase pairs, runs alongside the patient and control lanes and is used to determine the mass (size) of any DNA fragments in the patient and control samples (Figure 31-13). Fluorescent dyes such as ethidium bromide and Gel Red® (which intercalate between the base pairs of the DNA helix) or SYBR green® (which binds to the minor groove of the DNA helix) are employed to visualize the DNA fragments of the patient, controls, and size markers in the gel. The newer dyes (e.g., Gel Red and SYBR green) have largely replaced ethidium bromide because they are much less toxic. Gels are soaked in a solution of diluted dye and then exposed to UV light, which causes the nucleic acid to appear as fluorescent bands. The mass of the bands in the patient and control lanes is determined by comparing the distance they migrated in the gel with the distance migrated by the bands of the size markers. Gel electrophoresis is appropriate when the goal is qualitative—that is, to determine the presence or absence of the target DNA.


FIGURE 31-13 Electrophoresis pattern of a DNA sample on a slab gel. A, Molecular size marker (ladder); B, positive control; C, negative control; D, no-DNA control. DNA samples are placed in wells at the cathode (negative pole) and migrate to the anode (positive pole) due to the negative charge of the DNA molecules. By comparing the bands present in the gel with the molecular size markers, the mass of each band, measured in base pairs, is determined. For example, in the positive control sample, the three bands are 184, 110, and 89 bp. Positive, negative, and no-DNA controls must be used when performing gel electrophoresis. The positive control contains the target DNA sequence, and the negative control lacks this sequence. The no-DNA control sample lacks DNA. No banding should be present in the no-DNA control. If bands are present, contamination of samples occurred during the testing process.

Another method of fractionating DNA fragments by mass (size) is capillary gel electrophoresis. In this type of electrophoresis, long fused silica capillaries filled with derivatized acrylamide polymer are used for separation of single-stranded negatively charged DNA fragments on the basis of size or number of base pairs. The sample is applied to the capillary using electrokinetic injection, and the DNA fragments are separated using a high voltage as they migrate through the capillary from the negative electrode to the positive electrode. Smaller DNA fragments move faster through the polymer in the capillary compared to larger fragments. Detection of the separated fragments occurs by incorporating a fluorescent label into the PCR-amplified DNA. Before reaching the positive electrode the fluorescently labeled DNA fragments cross the path of a laser beam and detector. When the laser beam hits a fluorescent DNA fragment, light is emitted at a specific wavelength. The light emission is read by the detector, and the signal produces a peak on an electropherogram (). Figure 31-14


FIGURE 31-14 An electropherogram of capillary gel electrophoresis showing the separation of fluorescently labeled DNA fragments by size (number of base pairs). Fragments migrate through the matrix in the capillary from the negative to the positive pole and emerge from the capillary in size order (smaller fragments first). Before reaching the positive electrode, the fluorescently labeled DNA fragments are detected by passing one at a time between a laser beam and the detector. Each fragment is represented by a peak on the electropherogram.

Capillary electrophoresis offers a number of advantages over traditional gel electrophoresis. The injection, separation, and detection of the fragments are automated. The separation can be quite rapid with excellent resolution. The time of fragment elution and the peak height information are stored for easy retrieval. Size ladders can be labeled with different fluorescent dyes and run in the same capillary as the sample providing more accurate sizing.31 This method of separation is used in a number of applications including B and T cell clonality testing, bone marrow engraftment analysis, and screening for the internal tandem duplication mutation in the FLT3 gene in acute myeloid leukemia (AML) (Chapter 35).32

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 allow recognition of many sequences. The number of restriction fragments produced depends on the number of restriction sites present in the amplified target.3334 Enzyme action at one restriction site produces two restriction fragments, action at two restriction sites produce three restriction fragments, and so on.

A restriction enzyme detects even a single base substitution if the mutation alters the recognition sequence and prevents digestion at the site or creates a new site resulting in an additional fragment. Arestriction fragment length polymorphism (RFLP) is a mutation or polymorphism-induced change in the recognition site of the restriction enzyme that alters the length (number of base pairs) of the restriction fragment. A mutation in the coagulation factor V gene (F5 Leiden mutation) is an excellent example of RFLP. Individuals possessing this mutation have an increased risk of venous thrombosis. This mutation results from the replacement of guanine with adenine at nucleotide position 1691 (G1691A) of the F5 gene.3536 The mutation alters a restriction site normally detected and cut by the restriction enzyme Mnl I. The wild-type (normal) F5 amplicon is 223 bp long with two Mnl I-specific sites. After PCR amplification and incubation with Mnl I, the wild-type amplicon is cut into three restriction fragments, separable using slab gel or capillary electrophoresis. 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 expected fragments 37, 82, and 104 bp. A sample from an individual homozygous for the F5 Leiden mutation possesses two copies of the mutated F5 gene and generates only two bands: 82 and 141 bp. A sample from a heterozygous individual possesses one normal and one mutated F5 gene and produces four bands of lengths 37, 82, 104, and 141 bp (Figure 31-15 and Figure 31-1).


FIGURE 31-15 Diagram of the amplified target sequence of the coagulation factor V (F5) gene (223 bp) in a PCR-Restriction Fragment Length Polymorphism (RFLP) method to detect the F5Leiden mutation. A, The amplified target sequence for the normal (wild-type) F5 gene contains two restriction sites for Mnl I, so three restriction fragments of 37, 82, and 104 bp are produced. B, In the F5 Leiden mutation, the substitution of an A for a G in position 1691 of the F5 gene eliminates one of the restriction sites for Mnl I. Thus the mutated F5 Leiden gene possesses only one restriction site for Mnl I. In individuals homozygous for the mutation, only two restriction fragments of 82 and 141 bp are produced. C, An individual who is heterozygous for the F5 Leiden mutation has a normal and a mutated F5 gene. Mnl I produces four restriction fragments of 37, 82, 104, and 141 bp.

Nucleic acid hybridization and southern blotting

Once used in a number of molecular tests, including B and T cell clonality assays, as well as the detection of chromosome translocations, Southern blots are now largely used for samples that do not provide a result using standard PCR methods or for research applications. Southern blots can only be performed on high-quality genomic DNA or PCR amplicons. Briefly, DNA is digested with a restriction enzyme, size fractionated using agarose gel electrophoresis, denatured to become single stranded, and then finally transferred to a solid support—typically a nylon or nitrocellulose membrane and then detected with a labeled probe.

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, unhybridized probe is washed off, and the hybridized bands are visualized, depending on the labeling system chosen. Most probes today are detected using a chemiluminescence detection by autoradiography ().Figure 31-1637-41


FIGURE 31-16 Southern blot steps. 1, DNA is cut with the restriction endonuclease EcoRI, which produces many restriction fragments. 2, DNA fragments are separated on an agarose gel. 3,DNA fragments are transferred to a nitrocellulose filter. 4, A labeled probe is hybridized to the DNA fragments on the filter. 5, Autoradiography is used to visualize the hybridized DNA probe, the detection of which indicates the presence of the given DNA sequence in the sample.

Cleavage-based signal amplification

Cleavage-based amplification is an isothermal signal amplification method marketed as Invader® (Hologic, Inc., Bedford, MA). In the primary reaction, the 3′ end of a test probe and an Invader oligo probe anneal to complementary sequences on the target DNA template forming a specific substrate site recognized by a Cleavase enzyme (Figure 31-17). The 5′ end of the test probe (5′ flap) does not anneal to the target. The Cleavase enzyme cuts and releases the 5′ flap of the test probe. In a coupled secondary reaction, the 5′ flap of the test probe anneals to a complementary signal probe that has a FRET (fluorescence resonance energy transfer) reporter. The signal probe and reporter (called a FRET cassette) is specific for the 5′ flap of the test probe. The FRET reporter is a fluorescent dye bound to the signal probe in close proximity to a quencher; the quencher prevents the reporter dye from emitting a fluorescent signal. The combination of the 5′ flap of the test probe and the FRET cassette forms another specific substrate site for the Cleavage enzyme. Cleavage of the 5′ end of the FRET cassette results in separation of the fluorescent reporter from the quencher and production of a fluorescent signal. Repeated binding and cleavage result in the signal amplification. Two reactions are done simultaneously using different fluorescent molecules for detection of either the wild-type or mutant sequence. Because the mutant sequence is different from the wild-type sequence, the wild-type test probe will not anneal to the mutant target, and no fluorescent signal will occur. This technique can be used to detect single base pair changes, small insertions, and deletions. It is also FDA-approved for the detection of mutations associated with thrombophilia, including the factor V (F5) Leiden mutation, the prothrombin G20210A mutation in the F2 gene, and methylenetetrahydrofolate reductase (MTHFR) gene mutations.42


FIGURE 31-17 Cleavage-based signal amplification. The Cleavage-based DNA Signal Amplification Assays use a Cleavase® enzyme to recognize and cleave specific structures formed by the addition of two oligonucleotide probes to a nucleic acid target. Two oligonucleotide probes (a test probe and an Invader® oligo probe) hybridize in tandem to the target DNA to form an overlapping structure. The 5′ end of the test probe includes a 5′ flap that does not hybridize to the target DNA. The 3′ nucleotide of the bound Invader® oligo overlaps the test probe. The Cleavase® enzyme recognizes this overlapping structure and cleaves off the unpaired 5′ flap of the test probe, releasing it as a target-specific product. In the secondary reaction, each released 5′ flap anneals to a fluorescence resonance energy transfer (FRET™) probe to create another overlapping structure that is recognized and cleaved by the Cleavase® enzyme. When the FRET™ probe is cleaved, the fluorophore (F) and quencher (Q) on the FRET™ probe are separated, generating detectable fluorescence signal. The initial and secondary reactions run concurrently in the same well. Two different fluorescent labels are used, one to identify the presence of the wild-type allele and one to identify the presence of the mutant allele.

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. A combination of these two important techniques (cycle sequencing) has made DNA sequencing an integral part of molecular diagnostics. In cycle sequencing, the order of the nucleotide bases is determined after amplification.43 Cycle sequencing is applied in molecular testing to assess amplified sequences for insertions, deletions, or point mutations, such as the FLT3 internal tandem duplication (ITD) or point mutations in the KIT gene that occur in AML (Chapter 35).44-46

Cycle sequencing is based on dideoxynucleotide terminator sequencing.47 The addition of nucleotides to a growing DNA 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 (Figure 31-5). If a nucleotide lacks the 3′ hydroxyl group, it can be incorporated into the newly synthesized strand of the DNA but cannot be extended, so the fragment terminates at the “defective” base. If low concentrations of the terminators, dideoxyadenosine triphosphate, dideoxycytosine triphosphate, dideoxyguanine triphosphate, and dideoxythymine triphosphate, are included in the single primer PCR master mix used for sequencing, over a number of cycles a series of DNA fragments is produced that terminate at each successive base with each fragment differing in length by one nucleotide. This is called a ladder or nested series of fragments.

In the dye terminator method each of the four dideoxynucleotides in the PCR reaction is labeled with a different fluorescent dye so that each DNA fragment terminates in a labeled dideoxynucleotide corresponding to the sequence of the target DNA (, Figure 31-18 A). The specific fluorescent color of the DNA fragment identifies the terminal nucleotide. Alternatively, in the dye primer method, the primers are labeled with four different fluorescent dyes (corresponding to each nucleotide), and in separate tubes, each labeled primer is subjected to PCR with unlabeled dideoxynucleotides (Figure 31-18B). As in the dye terminator method, the specific fluorescent color of the DNA fragments corresponds to the terminal nucleotide.




FIGURE 31-18 Dideoxy Chain Termination (Sanger) DNA Sequencing. Cycle sequencing of a DNA template produces a nested series of fragments that differ by one nucleotide each. The template is amplified by polymerase chain reaction (PCR) using a single primer sequence (single-sided PCR). The PCR master mix includes small amounts of dideoxynucleotides. When a dideoxynucleotide is incorporated into the growing DNA polymer, chain extension terminates. A, Dye terminator method in which each of the four dideoxynucleotides are labeled with a different fluorescent dye. The identity of the terminal nucleotide corresponds to its specific fluorescent color. B, Dye primer method in which the primer is labeled with four different fluorescent dyes, corresponding to each nucleotide. Again, the identity of the terminal nucleotide corresponds to the specific color of the primer. C, Capillary gel electrophoresis in which fluorescently labeled fragments pass between a laser and detector in size order (smallest fragments first). The fluorescent color of the fragment identifies the terminal nucleotide. The signals appear as peaks on an electropherogram, with each peak representing a specific terminal nucleotide in sequence order.

The fluorescently labeled fragments are subjected to capillary electrophoresis (described earlier in the chapter). The DNA fragments migrate through the capillary and separate based on their size. Near the end of the capillary, the fragments pass one by one through the beam of a laser in an order based on their length (with the shortest fragments emerging first). A detector reads the specific fluorescent color of each fragment and displays the signal as a peak on an electropherogram; this allows the sequence to be read (Figure 31-18C).

In order to unambiguously read the nucleotide at each position, the PCR reaction for cycle sequencing contains only a single primer that produces single-sided PCR. Two separate reactions are typically carried out, one using the forward primer and a second using the reverse primer. This produces complementary sequences from both strands. After the cycle sequencing reactions, the nested products are purified and denatured before loading on the capillary sequencer. The injection, separation, and detection are automated, but the operator can set the parameters such as amount of sample injected, length of capillaries, and the type of polymer. Capillary DNA sequencing instruments are equipped with base calling software that will read the base sequence of the DNA fragment sequenced. Software packages will also identify alterations in the sequence such as single nucleotide polymorphisms (SNPs), point mutations, and insertions or deletions based on comparison to a specific reference sequence.

Pyrosequencing is another sequencing method that is useful for the determination of point mutations and short sequence analysis. This method uses a “sequencing by synthesis” principle and the detection of pyrophosphate release upon nucleotide incorporation. Nucleotides are added sequentially to a single-stranded template, and when the complementary base is added, it is incorporated, resulting in pyrophosphate release. The pyrophosphate released is then converted to ATP by sulfurylase in the presence of adenosine 5′ phosphosulfate. This reaction is coupled to the luminescent conversion of luciferin to oxyluciferin by the enzyme luciferase, resulting in the release of light. Luminescent reactions resulting from the incorporation of nucleotides are represented as peaks on a pyrogram. The intensity of the light determines if there are multiple nucleotides that are identical because the peak height in the pyrogram will be proportional to the number of nucleotides.48

Sanger DNA sequencing is considered the gold standard for the detection of point mutations and single nucleotide polymorphisms. With a point mutation, for example, sequencing of either strand will show whether the mutation (for example, adenine to thymine) is present by comparison of the sequence to the reference sequence. 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 a homozygote, the two nested series of fragments will be identical, whether wild-type or mutant. If the patient is a heterozygote, 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 signals will be present at the position of the mutation, but half the templates will contain each sequence.

Next-generation sequencing (NGS), also known as massively parallel sequencing (MPS), is a new technique that is being increasingly applied in all areas of molecular diagnostics, including hematology. The technology is still rapidly changing, but most of the currently available methods sequence short fragments multiple times and use bioinformatics to reassemble the sequence. The process of NGS can be divided into several steps, including template preparation, sequencing and detection, and finally data analysis and assembly. Currently available commercial systems use a variety of methods. One commonly used method involves the immobilization of molecules on a solid phase followed by amplification to produce clonally amplified clusters. Sequencing by synthesis reactions are carried out using cyclic reversible terminators in four colors and fluorescent detection by lasers following each base addition. A second commonly used method also amplifies the sequencing template but uses emulsion PCR to accomplish it. The sequencing technology takes advantage of the hydrogen ion released when a base is added and uses semiconductor technology to translate that into a nucleotide sequence by the sequential addition of bases and the measurement of the voltage produced when the correct nucleotide base is added. Both methods use proprietary software and alignment to a reference sequence to produce the final template sequence. There are also numerous programs available as open source or from commercial vendors for analysis. Current applications for NGS have been mainly limited to the sequencing of panels of genes associated with a particular disease. This makes the bioinformatics analysis more manageable and limits the number of variants of unknown significance (VUS) that are identified.49-51

Real-time polymerase chain reaction

In contrast to end-point PCR, real-time PCR measures the change in nucleic acid amplification as replication progresses using fluorescent marker dyes. There are several commercially available instruments that vary in their capacity, sample volume, and optics.52 There are a variety of choices in the optics for fluorescent detection. A tungsten lamp is commonly used for excitation, and different filters are used to select the excitation and emission wavelength. Light emitting diodes (LEDs) or lasers for excitation can also be coupled with emission detection, depending on the instrument. Real-time PCR can be used in quantitative or qualitative assays. The time interval, expressed as the number of replication cycles, required to reach a selected fluorescence threshold is proportional to the copy number of target molecules in the original sample.53The PCR cycle at which amplification crosses the threshold is denoted as the Ct for threshold value or the Cp for crossing point value. Importantly these values are calculated from the exponential portion of the amplification curve. The Ct value is inversely related to the amount of target so that the more starting DNA or cDNA that is present in the reaction, the lower the number of PCR cycles that are required to reach the threshold and exponential phase of the reaction (Figure 31-19).


FIGURE 31-19 Real-time PCR amplification curve illustrating the important features of the curve, including the threshold CT value and the exponential phase of the curve.

Real-time PCR requires the use of fluorescent detection, and there are several different options available. The simplest and the most straightforward option is to add a fluorescent dye, such as SYBR® green, to the PCR reaction. These dyes bind to double-stranded DNA so that the fluorescence increases in proportion to the number of copies of the PCR product. The disadvantage of this approach is that these dyes do not differentiate between specific and nonspecific PCR amplicons, so the PCR reaction must be free of mispriming and primer dimers (primers that partially anneal to one another and are extended by the polymerase, forming very short amplicons). A more specific method of detection adds a probe in addition to the forward and reverse PCR primers, which also binds to the amplicon, providing additional specificity. There are several methods commonly employed, including hybridization probes, Taqman probes, and molecular beacon and Scorpion probes. Hybridization probes utilize two oligonucleotide probes that bind to the amplicon adjacent (within one to five bases) to one another. One of the oligos has a 3′ donor fluorophore and the other a 5′ acceptor. The 3′ fluorophore is excited, and the energy is transferred to the acceptor, which then fluorescences at a detectable wavelength. This is called fluorescence resonance energy transfer, or FRET.54 Hybridization probe technology in combination with melting curve analysis is used in some commercial thrombophilia assays.

Taqman probes consist of a single oligonucleotide that anneals between the forward and reverse primers. This probe contains a fluorophore on the 5′ end and a quencher on the 3′ end. This method takes advantage of the 5′-to-3′ exonuclease activity of DNA polymerase. As the amplicon is synthesized by DNA polymerase, the probe is degraded. This separates the fluorophore from the quencher and results in fluorescence. As the number of amplicons increases, there is more target to anneal to the probe and greater fluorescence as the probes are degraded.55 Molecular beacons and Scorpion probes use a hairpin structure to juxtapose the reporter and quencher. When the probe binds to the target, the hairpin unfolds and separates the fluorophore from the quencher, and fluorescence is detected.56

Qualitative real-time polymerase chain reaction

Taqman assays are used to detect point mutations such as the common point mutations in hereditary hemochromatosis (Chapter 20). Two Taqman probes are synthesized with a different fluorescent label, complementary to either the wild-type or mutant sequence. If the sequence is complementary to the target, the probe will be degraded as described above, and fluorescence will be produced. If the sequence contains a mismatch, the probe will be displaced and the fluorescence will remain quenched. Real-time PCR can also be combined with sequence- specific primer PCR (SSP-PCR) to detect point mutations or SNPs. This method takes advantage of the fact that the 3′ end of a primer in PCR must match the template sequence exactly to be extended by the polymerase. This is in contrast to the 5′ end of the primer, which can have additional nucleotides added. By using primers complementary to either the wild-type or mutant nucleotide, the presence or absence of a mutation can be determined by the reaction that produces the PCR product.57 There are several modifications of this technique. One widely used technique, called SNaPshot®, uses dideoxynucleotides, each labeled with a different fluorophore and primers of different size in a multiplex reaction to detect several different nucleotide changes simultaneously. The different-sized PCR products are then identified by size fractionation using capillary gel electrophoresis (Figure 31-20).58There are other variations on this technique as well that are not discussed here. It is clear, however, that there are an increasing number of applications of real-time PCR in hematology, and multiple different techniques can be used to detect the same mutation. Applications of these techniques include the detection of resistance mutations in viruses or bacteria, as well as somatic mutations in cancer cells and germline mutations in genetic diseases.59



FIGURE 31-20 Qualitative real-time PCR for hereditary hemochromatosis, HFE C282 gene mutation detection. The mutation replaces the amino acid cysteine (C) with tyrosine (Y) in position 282 of the HFE protein. The method uses two Taqman amplification probes, one for the wild-type (normal) and one for the mutant allele, each labeled with a different fluorescent reporter. A, Taqman allelic discrimination plot demonstrating the three genotypic populations for the HFE gene and no template controls. The scatter plots are derived from the total fluorescence of the amplification curve for both fluorescent probes. The genotype can be determined from the position on the scatter plot. B, Real-time PCR amplification curves of a heterozygous C282C/Y mutant patient showing fluorescence with the two Taqman probes: VIC for the wild-type and FAM for the mutant allele.

Quantitative real-time polymerase chain reaction

Real-time quantitative PCR can be done in two ways: relative quantitation, which normalizes to a reference gene used when measuring gene expression, or absolute quantitation, where a standard curve of known copy number of diluted standards is run along with the patient samples. Once the relationship between the copy number of the standards and the Ct value is determined using the standard curve, the copy number of the patient sample can be determined from the crossing point value. This is used to monitor residual disease in chronic myelogenous leukemia by quantifying the amount of the BCR-ABL1 transcript (Chapter 33) as well as viral loads in infectious disease (Figure 31-21).60-66


FIGURE 31-21 Quantitative real-time PCR for BCR-ABL1 transcript showing the standard curve with standards containing known copy numbers of the BCR-ABL1 cDNA. Plotting the crossing point threshold value (CT) on the y-axis versus the log of the copy number on the x-axis generates a standard curve.

High Resolution Melting Curve (HRM) analysis is a real-time PCR method that uses the quantitative analysis of the melting curve to detect sequence differences in PCR amplicons. Melting curves are often run in conjunction with real-time PCRs to confirm specificity of the product by heating the amplicon in increasing intervals from 65° C to 95° C and measuring the fluorescence. When the double-stranded DNA melts, the fluorescence will sharply decrease. High-resolution melt curves use narrower temperature increments and a saturating fluorescent dye to determine the melting curve. This allows the determination of sequence differences in PCR amplicons. This method requires a thermocycler with good temperature stability and a software package for HRM analysis. The advantage of HRM is that it will detect any sequence difference in an amplicon and the exact mutation does not have to be known in advance.67

Minimal residual disease in leukemia

Real-time quantitative PCR provides the opportunity to follow disease burden and to measure minimal residual disease (MRD) in leukemia, a key indicator of treatment efficacy, clinical remission, and prognosis.6162Currently, chemotherapy, radiation therapy, and hematopoietic stem cell transplantation reduce leukemic cells to levels undetectable first by visual examination of a bone marrow smear or peripheral blood film and later by flow cytometry assay.68 The persistence of disease after treatment that is only detected by molecular assays is called minimal residual disease. Real-time quantitative PCR identifies the specific nucleic acid sequence in residual leukemic cells and helps guide the types and intensity of therapy with the goal of “molecular” remission. Real-time reverse transcriptase quantitative PCR to assess the fusion transcript levels for CML is regarded as the “gold standard” for the detection and quantification of minimal residual disease. Subsequent to remission, periodic real-time quantitative PCR assays are used to detect early relapse and drug resistance, enabling the hematologist to initiate appropriate follow-up therapy.69

Real-time quantitative PCR may detect a few malignant cells within a population of a million cells, providing unparalleled sensitivity. Current assays to assess MRD can detect one leukemic cell among 105 to 106normal cells. Current applications include detection of BCR-ABL1 transcripts in CML and some acute leukemias (Figure 31-22); JAK2 (“just another kinase” or Janus kinase) mutations in the myeloproliferative neoplasms, polycythemia vera, and essential thrombocythemia (Chapter 33); the t(15; 17)(q22; q21) or PML-RARA fusion transcript in acute promyelocytic leukemia (Chapter 35); and gene rearrangements in mature lymphoid neoplasms (Chapter 36).70


FIGURE 31-22 Quantitative real-time PCR for BCR-ABL1 transcript. Using the standard curve, the copy number of BCR-ABL1 transcripts in unknown samples can be determined. Two amplification graphs are shown. The graph on the left shows the amplification of the BCR-ABL1 transcript, and the graph on the right shows the amplification of the ABL1 transcript as a control for sample quality.

A major issue with quantitative assays has been the lack of reproducibility between different laboratories due to the specimen type and quality, the choice of housekeeping gene for normalization, and the specific assay used. Recently an international standard for BCR-ABL1 has been developed and made available by the World Health Organization (WHO). This standard will serve as a universal standard and allow for interlaboratory comparison.71

Mutation enrichment strategies

In order to detect low levels of disease or emerging resistance, it is helpful to be able to enrich for the presence of the mutation. There are currently several methods to accomplish this, all of which seek to selectively amplify the mutant sequence in the presence of an excess of wild-type sequence. Peptide-nucleic acid (PNA) and locked nucleic acid (LNA) both contain normal nucleotide bases for hybridization but different backbones from the phosphodiester backbone of DNA and RNA.72 This gives these probes the ability to hybridize more tightly when used as probes in PCR reactions. When the probes span and match the wild-type sequence, they can inhibit the amplification of the wild-type allele, thus enriching for the mutant allele.73

COLD-PCR (co-amplification at lower denaturation temperature–PCR) is another mutation enrichment technique based on PCR amplification. COLD-PCR is based on the principle that DNA containing a mismatch will melt at a slightly lower temperature than completely matched sequences. Designing the PCR cycle temperatures to maximize that difference results in a preferential amplification of the mutant sequence in a mixed sample of mutant and wild-type DNA, even when the mutant is in very low concentrations. This is accomplished by carrying out the denaturation step at the temperature that will have mutant-wild-type heteroduplexes in a single-stranded state, while wild-type homoduplexes will not yet have denatured.74 All of these methods, although useful to enrich for mutant alleles, are technically demanding and therefore are not yet in widespread usage in molecular laboratories.

Chromosome microarrays

Chromosomal microarray analysis is a methodology used to measure gains and losses of genomic DNA. The advantage of microarrays compared to karyotyping is that it is a higher-resolution method and will detect genetic changes that cannot be observed by karyotyping (Chapter 30). In addition, chromosome microarrays have the advantage of also detecting aneuploidy and large chromosomal duplications and insertions.

There are two different types of chromosome microarrays: comparative genomic hybridization (aCGH) and single nucleotide polymorphism array (SNP-A) karyotyping. Both types of arrays can identify variation in copy number. Due to differences in methodology, however, they detect different types of variants (). Figure 31-23



FIGURE 31-23 Chromosomal microarrays. A, Diagram of the procedure for microarray-based comparative genomic hybridization (aCGH). B, An example of a single nucleotide polymorphism array (SNP-A) karyogram from a patient with a secondary acute myeloid leukemia with microdeletions on chromosomes 4 and 19 and normal chromosome 9. Source: (© 2008 SLACK, Inc. Modified from Shaffer L. G. & Bejjani B. A. Using microarray-based molecular cytogenetic methods to identify chromosome abnormalities. Pediatric Annals 38, 440-447 (2009) doi: 10.3928/00904481-20090723-08.)

In an array-based assay, the specimen DNA is isolated, denatured, and hybridized to a chip or array containing thousands of probes with known sequences. For comparative genome hybridization, the patient and a control DNA are labeled with different fluorescent dyes, and after hybridization, the relative intensity of the two fluorescent signals is used to determine if there are any genomic gains or losses. Duplications result in a higher-intensity fluorescence relative to control, and deletions result in a lower-intensity fluorescence. CGH is most useful in the detection of relatively large duplications or deletions.75

In SNP arrays only the patient DNA is labeled, denatured, and hybridized to an array containing probes with known SNPs. Again, the signal intensity is used to determine the copy number. SNP arrays are able to detect runs of homozygosity that can indicate uniparental disomy or consanguinity.76 Because each of these methods has both advantages and disadvantages, array platforms have been developed that contain both types of sequences: SNPs and larger clones used in CGH. This provides a more uniform coverage over the entire genome. In certain situations, arrays are replacing or used as an adjunct to conventional karyotyping and fluorescence in situ hybridization (FISH) (Chapter 30). Arrays are useful to detect copy number variants but do not detect balanced translocations.77-78

Pathogen detection and infectious disease load

Box 31-4 contains a listing of hematologically important pathogens detected by molecular methods. 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.79 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 rapid 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 effective 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. There are currently multiple FDA approved assays for the detection of viral and bacterial pathogens. A current listing of these tests can be found in the test directory on the Association for Molecular Pathology web site (

Clinical relevance is important when assessing infectious disease using molecular techniques. These methods allow millions of copies to be generated from a single DNA or RNA 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. 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.

Current developments

Molecular diagnostics is a rapidly growing area of the clinical laboratory, and the technology continues to develop. It promises to revolutionize laboratory techniques in all disciplines, and the technologies of genomics are being extended to proteomics (the molecular analysis of proteins) and metabolomics (the molecular analysis of metabolism). Methods continue to be automated and miniaturized, providing ever greater sensitivity and reliability coupled with short turnaround time and technical simplification. In many situations, assays are moving from single analyte assays to multiplex assays detecting panels of analytes. In the case of leukemias such as AML, mutations in multiple genes are incorporated into the WHO guidelines.80 Methods such as the SNaPshot technique described previously are being applied to detect multiple mutations simultaneously.

Another technique being applied to the detection of mutation panels is Matrix-assisted laser desorption/ionization—time of flight (MALDI-TOF) mass spectrometry. This methodology uses PCR coupled to a single-base extension reaction that adds labeled nucleotides so that the extension products containing different mutations have different masses. These reactions are also multiplexed to increase throughput, detecting hundreds of mutations in a single panel assay.81

Digital PCR (ddPCR) is a technique with very high sensitivity that can be used to detect resistance mutations to tyrosine kinase inhibitors used to treat CML (such as the T315I resistance mutation in the BCR-ABL1gene)82 (Chapter 33) or to quantify virus copy number.83 This technique uses various methods—for example, a droplet generator to create nanoliter droplets that partition template molecules, which are then amplified by PCR. The amplicons are detected by fluorescence and either read by a droplet reader or other detection mechanism. For translocation detection, wells containing an amplified housekeeping gene, translocation product, or both are then quantitated. This method is extremely sensitive, detecting a few molecules per sample, and can be applied to both DNA and RNA applications.

NGS will continue to play an increasingly important role in molecular diagnostics. In addition to sequencing panels of genes, this technology has been used to sequence whole genomes, exomes (the coding exons), as well as RNA sequencing (RNAseq)8485 (Figure 31-24). This technology is also being applied to the determination of the epigenome86—modifications such as methylation that affect gene regulation and expression.




FIGURE 31-24 A, Diagram of the procedure for next-generation sequencing (NGS) using the two most common technologies. B, An illustration of a number of NGS reads for a 32-nucleotide sequence aligned with the genomic reference sequence in blue on the bottom (sequence mismatches are in red). The center of the alignment shows a variant present in the heterozygous state. C, Sequenced fragments are depicted as bars with colored tips representing the sequenced ends and the unsequenced portion of the fragment in gray. Reads are aligned to the reference genome (for example, mostly chromosome 1 in this example). The colors of the sequenced ends show where they align. Different types of genomic alterations can be detected. From left to right, point mutations (in this example, A to C) and small insertions and deletions (indels) (in this example, a deletion shown by a dashed line) are detected by identifying multiple reads that show nonreference sequence; changes in sequencing depth (relative to a normal control) are used to identify copy number changes (shaded boxes represent absent or decreased reads in a tumor sample); paired-ends that map to different genomic loci (in this case, chromosome 5) are evidence of rearrangements; and sequences that map to nonhuman sequences are evidence for the potential presence of genomic material from pathogens. Source: (A from Grada A, Weinbrecht K: Next-generation sequencing: methodology and application. Journal of Investigative Dermatology 133, 248-251, 2013; Bfrom Almomani R, van der Heijden J, Ariyurek Y, et al: Experiences with array-based sequence capture; toward clinical applications. European Journal of Human Genetics 19:50-55, 2011; C from Meyerson M, Gabriel S, Getz G: Advances in understanding cancer genomes through second generation sequencing, Nature Reviews Genetics 11: 685-696, 2010.)

Small microRNAs (miR; ∼20 nucleotides long) that were once thought to be insignificant are evolving as biomarkers for the progression of hematologic malignancies. The dysregulation of miRs affects normal hematopoiesis, and their atypical expression is beginning to be established in T and B cell leukemias and lymphomas.87 These miRs target genes in the 3′ UTR (3′ untranslated region) and are hypothesized to inhibit the translation of mRNA to proteins.88 These miRs can be detected by many molecular-based techniques such as PCR, NGS, and microarray technology. Their clinical role in hematologic cancer therapy as a prognostic marker is now starting to be recognized. A link between p53 expression and possible therapeutic efficacy is shown with miR-181a/b dysregulation in chronic lymphocytic leukemia (CLL) patients in a recent study.89 Another example is a report by Seca and colleagues90 that lists many functions of miR-21, including the upregulation of the BCR-ABL1 protein and induction of chemoresistance. An overexpression of miR-21 has been demonstrated in patients who are fludarabine nonresponders.91 The overexpression of plasma miR-155 is correlated to the identification of B-CLL in patients,92 and association of miR-21 is established with drug resistance in plasma cell myeloma.93 The listed studies demonstrate that the ability to measure the expression of miR has expanded the repertoire of diagnostic, prognostic, and therapeutic efficacy markers in hematologic malignancies.

The future molecular technologies will increase the efficiency and sensitivity for detection of all types of genome alterations, including point mutations, insertion and deletion mutations, copy number variants, and chromosome rearrangements. It will facilitate the discovery of new chromosome rearrangements as well as the diagnosis of microbial infections. It will result in refined classification and improved treatment of hematological diseases.


• DNA directs cell function as described by the central dogma. DNA retains the genetic code and reproduces itself through replication.

• The genetic code is transcribed from DNA to mRNA. mRNA consists of coding exons and noncoding introns that are excised after transcription. The processed mRNA then transports the code from the nucleus to the cytoplasm, where it is translated by the cytoplasmic ribosomes. Translation depends on tRNA, small RNA molecules designed to transport and add amino acid to growing peptide chains in cytoplasmic ribosomes.

• DNA consists of a five-carbon sugar (deoxyribose), a phosphate group, and a nitrogenous base. The bases are either purines or pyrimidines. The purines in DNA are adenine (A) and guanine (G). The pyrimidines in DNA are thymine (T) and cytosine (C).

• DNA is a double-stranded molecule held together by hydrogen bonding between the bases, A to T and G to C. Heat denatures double-stranded DNA by breaking the hydrogen bonds to produce single-stranded DNA. DNA strands are antiparallel.

• RNA is a single-stranded molecule that contains the sugar ribose instead of the deoxyribose found in DNA and the pyrimidine uracil in place of thymine.

• RNA polymerase recognizes a sequence of deoxyribonucleotides called the promoter within DNA. RNA polymerase separates the DNA strands and begins adding ribonucleotides, forming an initial RNA transcript consisting of introns and exons.

• Proteins function as structural components of the cell, as enzymes involved in metabolism or regulation, as receptors to regulate cellular functions, or as antibodies for the immune system. Mutation within a gene ultimately alters the protein produced, which often affects the function of the protein.

• Five areas of hematopathologic molecular testing include detection of mutations, gene rearrangements, and chromosomal abnormalities for diagnosis and prognosis of hematologic malignancies; detection and quantification of minimal residual disease to monitor treatment of hematologic malignancies; detection of mutations in inherited hematologic disorders; pharmacogenetic testing to detect genetic variation affecting certain drug therapies; and identification of hematologically important infectious diseases.

• Peripheral blood, bone marrow, tissue biopsy samples (both fresh and formalin fixed paraffin-embedded), fine-needle aspirates, body fluids, saliva, and cheek swabs are specimens used for DNA and RNA isolation.

• DNA is amplified by in vitro endpoint polymerase chain reaction (PCR) and real-time PCR. RNA targets can be amplified by PCR by first converting the RNA target to complementary DNA or cDNA.

• In endpoint PCR, amplified DNA is detected by electrophoresis (slab gel or automated capillary gel electrophoresis) or cleavage-based signal amplification (Invader) technology. Endpoint PCR can also be combined with restriction enzyme digestion of the amplicons, followed by detection of the restriction fragments by one of the methods mentioned above. In real-time PCR, the amplicons are detected during the PCR cycles by fluorescence detection.

• Real-time PCR can be used qualitatively to determine the presence or absence of a target or can quantify the copy number of a target DNA or RNA. PCR can be used to amplify RNA targets by first converting them to cDNA with reverse transcriptase activity.

• The dideoxy chain termination (Sanger) method for DNA sequencing is based on the principle that synthesis of a DNA polymer is terminated upon incorporation of a dideoxynucleotide. The target DNA template is amplified over a number of cycles, which produces a series of DNA fragments that terminate at each successive base with each fragment differing in length by one nucleotide. DNA fragments are detected by labeling either the dideoxynucleotide or the primer with a fluorescent dye. Other methods for DNA sequencing include pyrosequencing and next-generation sequencing (NGS).

• Chromosomal microarrays measure gains and losses of genomic DNA. They provide much greater sensitivity in detecting small genomic changes compared to conventional karyotyping. There are two types: microarray-based comparative genomic hybridization (aCGH) and single nucleotide polymorphism array (SNP-A) karyotyping. CGH requires the use of a control DNA.

• Molecular testing permits clinicians to make more accurate diagnostic, therapeutic, and prognostic decisions. It also allows a more sensitive assessment of minimal residual disease and therapeutic efficacy, resulting in better patient management.

Now that you have completed this chapter, go back and read again the case study at the beginning and respond to the questions presented.

Review questions

Answers can be found in the Appendix.

1. If the DNA nucleotide sequence is 5′-ATTAGC-3′, then the mRNA sequence transcribed from this template is:

a. 5′-GCUAAU-3′

b. 5′-AUUAGC-3′

c. 5′-TAATCG-3′

d. 5′-UAAUCG-3′

2. Cells with damaged DNA and mutated or nonfunctioning cell cycle regulatory proteins:

a. Are arrested in G1 and the DNA is repaired

b. Continue to divide, which leads to tumor progression

c. Divide normally, producing identical daughter cells

d. Go through apoptosis

3. To start DNA replication, DNA polymerase requires an available 3′ hydroxyl group found on the:

a. Leading strand

b. mRNA

c. Parent strand

d. Primer

4. Ligase joins Okazaki fragments of the:

a. 5′-to-3′ template strand

b. Lagging strand

c. Leading strand

d. Primer fragments

5. A 40-year-old patient enters the hospital with a rare form of cancer caused by faulty cell division regulation. This cancer localized in the patient’s spleen. An ambitious laboratory developed a molecular test to verify the type of cancer present. This molecular test would require patient samples taken from which two tissues?

a. Abnormal growths found on the skin and in the bone marrow

b. Normal splenic tissue and cancerous tissue

c. Cancerous tissue in spleen and bone marrow

d. Peripheral blood and cancerous tissue in the spleen

6. One main difference between PCR and reverse transcriptase PCR is that:

a. PCR requires primers

b. PCR uses reverse transcriptase to elongate the primers

c. Reverse transcriptase PCR forms millions of cDNA fragments

d. Reverse transcriptase PCR requires ligase to amplify the target DNA

7. Which one of the following statements about gel electrophoresis is false?

a. The gel is oriented in the chamber with the wells at the positive terminal.

b. A buffer solution is required to maintain the electrical current.

c. The matrix of a polyacrylamide gel is tighter than that of an agarose gel.

d. The larger DNA fragments will be closest to the wells of the gel.

8. Autoradiography of DNA is the:

a. Detection of radioactive or chemoluminescent oligonucleotides

b. Exposure of the gel to UV light

c. Transfer of DNA to a nitrocellulose filter

d. Use of ethidium bromide to visualize the DNA banding pattern

9. One major difference between endpoint PCR and real-time PCR is that:

a. Endpoint PCR requires thermostable DNA polymerase, deoxynucleotides, and primers

b. Endpoint PCR requires a separate step to detect the amplicons formed in the reaction

c. Real-time PCR uses capillary gel electrophoresis to detect amplicons during PCR cycling

d. Real-time PCR detects and quantifies amplicons using cleavage-based signal amplification

10. Which of the following statements about minimal residual disease is true?

a. Clinical remission of hematologic cancers is determined by molecular techniques such as PCR and flow cytometry.

b. Real-time quantitative PCR–determined copy number of BCR/ABL1 transcripts will always be lower in molecular remission than in clinical remission.

c. Qualitative PCR that uses a known copy number of a target sequence is of use in determining minimal residual disease levels.

d. Minimal residual disease assessment can aid physicians in making treatment decisions but does not yet offer insights into prognosis.


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*The authors acknowledge Dr. Mark E. Lasbury, whose work in prior editions formed the foundation for this chapter.