Template Generation

The general starting point for all NGS assays is double-stranded DNA (dsDNA). It is obtained from a variety of sources, including genomic DNA and reverse-transcribed RNA (i.e., complementary DNA [cDNA]). All starting dsDNA must be converted into a sequencing library, wherein fragmentation, size selection, and adapter ligation are used to generate an unbiased representation of the DNA population to be sequenced. Template generation spatially separates and immobilizes DNA fragments by attaching them to solid surfaces or beads. Limitations in the detection sensitivity require that sequencing reactions be amplified before sequencing, which may introduce bias (e.g. loss of representation of rare sequences) and errors (e.g., particularly problematic in clinical applications). The latest technologic platforms can sequence from single-molecule templates, which represents such a significant improvement that some authorities refer to them as third-generation platforms.⁴

Sequencing Reaction

NGS platforms use a series of repeating chemical reactions that apply DNA polymerase or ligase to add and detect nucleotides on a repetitive nucleotide-by-nucleotide basis. This process, referred to as sequence by synthesis, represents a dramatic improvement to Sanger sequencing, which requires discrete separation and detection of fragments that differ in length by 1 bp. Thee simultaneous analysis of massive numbers of reactions by NGS is largely responsible for the more than 100,000-fold decrease in per base sequence cost during the past 5 years.

Data Analysis

Compared with Sanger sequencing (which typically reads 500 bp to 1 kb in a single reaction), most NGS platforms offer shorter average read lengths (30 to 400 bp). Because NGS reactions generate millions of these short reads, data analysis is a much more difficult task, making heavy demands on data acquisition, storage, tracking, quality control, analysis, and interpretation.⁴ NGS data generation can vastly outpace analytic and interpretation resources.

Typically, the first phase of analysis is base calling, which is usually performed by the proprietary software associated with the sequencing platform. Base calling is followed by sequence alignment and assembly, an area of extremely active computational research.⁵ The last phase of analysis requires interpretation of the final sequence data. Third-generation platforms may offer longer reads, which could vastly reduce the challenges of data analysis, but these underdeveloped platforms are not widely available.

Sequencing Coverage

All NGS platforms have inherent errors in base calling, and the rate of error varies with each platform and chemistry. To counter these qualitative errors, each base pair is sequenced multiple times in separate, parallel sequencing reactions. The degree to which each nucleotide is quantitatively resequenced is referred to as the coverage or coverage depth of sequencing.

Because there may be biases in coverage depth during any NGS experiment, evolving standards suggest a need for 30 to 100 times coverage to ensure 100% accuracy in detection of single-nucleotide sequence variants. Deep sequencing produces very high coverage, meaning that each nucleotide may be sequenced hundreds to thousands of times, enabling the quantitative detection of rare variants in a DNA population. This method has clinical application in the detection of rare mutations. The occurrence of errors and the need for coverage contribute to the plateau in the cost reduction of whole-genome sequencing. This plateau may be overcome by the increased fidelity associated with third-generation technologies that use single-copy DNA.

Clinical Applications

NGS platforms offer the potential for large-scale genotyping and tumor profiling to characterize human disease at an individual level. Despite the explosion of research studies, validated clinical applications remain limited and are targeted at the small number of disease-associated genes. Targeted sequencing with NGS technology requires enriching DNA regions of interest before sequencing. Enrichment strategies include hybrid capture, microdroplet PCR, and array capture. Abnormalities identified in NGS screens are typically confirmed with traditional Sanger sequencing.

BEAMing Technology

BEAMing is a proprietary technology (Inostics, Baltimore, Md.) that allows the quantitative detection of oncogenic mutations.¹² The technology allows performance of single-molecule PCR on magnetic beads in an emulsion and can detect mutant DNA at low levels compared with wild-type DNA (ratios as low as 1 : 10,000). BEAMing has been used to detect tumor mutations in the circulating DNA and fecal DNA of colorectal cancer patients and to monitor the evolution of resistant KIT mutations in gastrointestinal stromal tumors (GIST) patients undergoing targeted therapies.^13,14

Detection of Minimal Residual Disease

The term “minimal residual disease” refers to persistence of neoplastic cells below the threshold of conventional morphologic detection. The rapid development of molecular technologies in the past decade has revolutionized the ability to monitor minimal residual disease. These molecular technologies have been particularly important to the management of acute myeloid leukemias and chronic myelogenous leukemia.¹⁵ Although the potential exists to apply these technologies to the management of many cancers, there have been no validated practical applications to date in management of GI cancers.

Fluorescence and Chromogenic In Situ Hybridization

FISH and chromogenic in situ hybridization (CISH) are widely used methods that can be applied to the detection of DNA and RNA. Metaphase FISH is a cytogenetic technique that requires culture of live cells and allows detection of much smaller abnormalities than are detected by conventional karyotyping. Interphase FISH does not require culture of live cells, and it is therefore a practical method that is commonly used to study routinely fixed tumors and hematologic malignancies.

For interphase FISH analyses, DNA probes in the 60- to 200-kb range are covalently attached to a fluorescent molecule, hybridized to a complementary target sequence in cellular DNA, and visualized under a fluorescent microscope as a point of fluorescent light in the nucleus of the cell. Probes are designed with specificity to any region of interest within the genome and are widely available for centromeric regions (i.e., CEN probes) and telomeres. Multiple probes may be hybridized simultaneously and analyzed separately by using different colors of fluorescent dyes.

Known, recurrent oncogenic translocations are detected by FISH by using dual-fusion probes or break-apart probes. Dual-fusion probes contain two probes, each labeled with a different fluorescent dye and designed to bind the regions spanning the breakpoint of each translocation partner. In the absence of a translocation, two distinct nuclear signals are observed for each colored probe. For a translocation, there is a single, distinct signal for each color from the nontranslocated chromosomes, and the two signals then combine both colors for the translocation and its reciprocal. Dual-color, break-apart probes are particularly useful in detecting translocations in which one chromosome can recombine with multiple partners. This approach consists of two probes that bind to the intact chromosome flanking the breakpoint. In the setting of a translocation, the two probes break apart from each other and yield two distinct signals rather than a single hybrid signal.

Copy number aberrations are well suited to detection by FISH. Centromeric probes give an accurate count of chromosome number and are useful in the detection of aneuploidy. When centromeric probes are used in combination with probes targeting hotspot chromosomal regions, they also allow a readout of regions of genomic deletion or amplification. Diploid tumor cells have two distinct centromeric probe signals and only one chromosomal arm signal in the setting of a deletion but more than two in the case of amplification. Multiple cells are typically scored, and a signal ratio is calculated and compared with validated positive and negative control ratios.

Polymerase Chain Reaction Detection of Translocations

Known, recurrent translocations are readily detectable by PCR. Different methods use the same general principles; several primer pairs and sets of primer pairs are designed to detect all possible known breakpoints for a given translocation. Multiple primers are needed because of the limitation in the size of PCR products and the large genomic variability in known breakpoints for some translocations. Assays usually are performed by real-time PCR with the use of standard curves for known controls, and results are reported as the percentage of translocated cells in the specimen tested.

Molecular Detection of Copy Number Aberrations

Copy number abnormalities may be detected by semiquantitative or quantitative PCR. For instance, PCR amplification of a polymorphic microsatellite reveals two bands in normal tissue. In the presence of an intragenic deletion in a tumor sample that spans the microsatellite, there is a reduction in amplification of the deleted allele. Referred to as loss of heterozygosity (LOH), this standard research approach characterizes genomic deletions in tumor samples and has been used in some clinical trials (e.g., evaluation of 18q LOH as a marker of adverse outcome in colorectal cancer). These types of assays are not being routinely used in validated tests in GI pathology.

Array Technologies

Array-based hybridizations were originally developed on nitrocellulose and nylon membranes and were moved to treated glass slides in 1987. The development of technology to deposit small spots of target on glass substrates led to a rapid increase in the miniaturization of spots and the array density of spots. The first automated arrayer was described in 1995, and it used a pen-type device to dot target material onto substrate.¹⁷ These technologies have rapidly evolved and are now capable of arraying as many as 100,000 spots on a substrate the equivalent size of a standard microscope slide.

Oligonucleotide arrays use technologies that directly synthesize DNA on glass or silicon substrates.¹⁸ These proprietary arrays (e.g., Affymetrix) use the efficient synthesis of short oligomers (10 to 25 bp), and samples can be arrayed at very high density. The selection of oligonucleotides to be arrayed varies with the intended application of the array.

Samples to be arrayed are typically prepared by fluorescent labeling, and several methods may be used. Some array technologies use single-color fluorescent labeling, whereas others (typically expression arrays) depend on dual-color competitive hybridizations in which a control sample is quantitatively compared with a test sample.

Reading microarrays requires a fluorescent reader and analysis software. Array results are corrected for background noise and normalized with standards. Results from duplicate or triplicate sample data are averaged by the analytic software.

Although very powerful in research settings, applications of array technology in the clinical molecular diagnostic laboratory remain limited, in large part because of a lack of established standards and controls and a high degree of noise and variability. The commercial development of many of these platforms has addressed many of these challenges, and array-based assays have recently seen increased use in clinical laboratories, particularly in the area of cytogenetics.

Expression Arrays

Expression arrays are designed to determine the relative expression level of a vast number of genes in a single sample. Typical experiments use labeled mRNA in a competitive, two-color fluorescent hybridization with a known control sample.¹⁹ These tests simultaneously measure the transcript level of thousands of genes compared with a control or normal specimen. This transcriptional profiling (i.e., transcriptome analysis) is commonly applied to human neoplasms.

Comparative Genomic Hybridization

CGH arrays are designed to test DNA. Specific genomic DNA sequences are spotted onto an array corresponding to loci known to be amplified or deleted in human tumors or possibly encompassing a much more comprehensive representation of the human genome.²⁰ Genomic DNA from the test sample is purified, fragmented, fluorescently labeled, and hybridized, typically in a competitive two-color hybridization with a known normal or control sample. To facilitate application to limited tissue samples, several methods have been developed to globally amplify test DNA before CGH analysis. Competitive hybridization allows a readout of the relative genomic copy number across all assayed genomic loci.

Universal Molecular Screening for Lynch Syndrome

Despite the identification of germline mismatch repair gene mutations as the cause of Lynch syndrome 20 years ago, identification and genetic testing of possible Lynch syndrome patients has been slow to catch on in clinical practice.²⁴ Although some of this disparity is related to a failure of the clinical community to rigorously screen all newly diagnosed colorectal cancer patients for a family history of cancer, many Lynch syndrome patients do not meet the traditional family history criteria. Approximately 40% of Lynch syndrome patients do not meet Amsterdam I or II criteria, including approximately 25% who do not meet the much less stringent Bethesda criteria.²⁵

Because effective screening tests are readily available and the morbidity and mortality associated with Lynch syndrome are greatly reduced with intensive colonoscopic screening, several authorities, including the American Molecular Pathology Association, recommend universal screening of newly diagnosed colorectal cancer by using microsatellite instability testing or immunohistochemistry.^26,27 Both tests have approximately 90% to 95% sensitivity for detection of a mismatch repair defect; in the setting of a strong family history, a second test is recommended if the initial screening test result is negative. Some universal screening programs use upper age cut-off values of 65 to 75 years. Figure 27-29 in Chapter 27 shows a sample screening algorithm that uses immunohistochemistry as an initial screening test. Immunohistochemistry offers several advantages as an initial screening test. It is rapid and inexpensive, does not require tissue microdissection or DNA extraction, can be readily preformed on a biopsy sample, and can be used to guide subsequent genetic testing in the case of an abnormality. Universal mismatch repair analysis is also useful for predicting patient outcomes and tailoring disease management.

Microsatellite Instability Testing

Microsatellite instability (MSI) testing is an accepted method of screening for Lynch syndrome, and MSI is the original molecular genetic signature by which the involvement of DNA mismatch repair defects in human cancer were first identified.²⁴ MSI testing is a PCR test that amplifies a panel of DNA microsatellites in tumor DNA and compares the results with normal DNA from the same patient. Because DNA mismatch repair deficiency is associated with a dramatic decrease in the fidelity of DNA repair, particularly at short repetitive DNA sequences, tumors with DNA mismatch repair deficiency have widespread alterations in the sizes of microsatellites, which are readily identified in an MSI test.

MSI testing is routinely performed on paraffin-embedded tissue and requires only crude tissue microdissection. After PCR amplification, amplified products are typically analyzed on an automated sequencing platform by capillary gel electrophoresis to resolve 1- and 2-bp shifts in microsatellite sizes (see Fig. 27-26 in Chapter 27). The use of relatively monomorphic mononucleotide repeats allows MSI testing to be performed on tumor DNA samples without accompanying normal DNA. A positive MSI test result has extremely high specificity for an underlying DNA mismatch repair deficiency.

Mismatch Repair Immunohistochemistry

The DNA mismatch repair protein complex is involved in postreplicative DNA repair and is therefore expressed in all proliferating cells, including normal crypt base epithelium and most colorectal cancers. Patients with Lynch syndrome inherit a mutant copy of one of the DNA mismatch repair genes, typically resulting in protein truncation or major impairment in protein function. Because the second allele is normally expressed in somatic tissues, there is no appreciable phenotype in normal cells. In tumorigenesis, the second wild-type allele is inactivated by point mutation, deletion, or methylation silencing, resulting in total loss of the normal mismatch repair function. Most inactivating mismatch repair gene defects result in total loss of mismatch repair protein expression, which is readily identifiable by immunohistochemisty.²⁴ The interpretation of mismatch repair protein immunohistochemistry is discussed further in Chapter 27.

Characterization of MLH1-Deficient Colorectal Cancers

Most MLH1-deficient colorectal cancers are caused by promoter hypermethylation of MLH1, which occurs in approximately 12% to 15% of colorectal cancers. The pathogenesis of methylation abnormalities in colorectal cancer is discussed in Chapter 27. MLH1 methylation silencing presents a distinct problem in the interpretation of MLH1 loss in universal screening programs; only approximately 15% of these cases are caused by Lynch syndrome.²⁴ Fortunately, two other extremely helpful tests can be used to determine which MLH1-deficient cases are likely to be caused by Lynch syndrome, and therefore need referral for genetic counseling and evaluation.

Germline Testing

Putative Lynch syndrome patients should be sent for genetic evaluation and counseling before germline testing, which is usually only undertaken only in patients whose tumors are mismatch repair deficient or have MSI. Rarely, tumors from Lynch syndrome patients have no detectable abnormalities by MSI testing and mismatch repair immunohistochemistry and some with normal screening results and a strong family history may still be referred for germline testing.

Genetic testing is performed by PCR and sequencing by using genomic DNA obtained from white cells from peripheral blood. Most mutations occur in MLH1 and MSH2 (see Table 23.3).²⁴ In addition to germline sequence variants and genomic deletions, rare families have heritable methylation silencing of MLH1 and MSH2 expression.^28,29

Immunohistochemical Analysis of HER2 Expression

Immunohistochemistry is the first test offered by most laboratories in the characterization of cancers for HER2, and the College of American Pathologists (CAP) guidelines state that algorithms should be accurate, reproducible, and performed in a CAP-accredited laboratory, thereby requiring rigorous validation and ongoing verification.⁵² Guidelines for scoring HER2 expression in gastric cancer are summarized in Table 23.4.^50,53 Briefly, the degree of lateral or basolateral membranous positivity is scored on a scale from 0 to 3+ (Fig. 23.2).

Fluorescence In Situ Hybridization Analysis of HER2 Copy Number

FISH is the gold standard for assessing the copy number of the HER2 gene. Dual-color, Food and Drug Administration (FDA)-approved commercial probe kits consist of a chromosome 17 centromeric reference probe and a 17q12 HER2 probe. Results are enumerated by counting 20 tumor cell nuclei and the ratio of the HER2 probe signals to the centromeric probe signals, with results higher than 2.2 considered positive for amplification (Fig. 23.3). FISH results may be scored manually or by using automated image analysis systems.

HER2 Testing in Other Cancers

In addition to breast and gastric cancer, HER2 is overexpressed in a subset of other cancers, including approximately 5% of colorectal cancers and some pancreatic, ovarian, endometrial, cervical, and lung cancers.^21,54 Clinical trials are ongoing for many of these cancers, and we may be entering an era in which widespread HER2 testing becomes a standard aspect of management of many advanced cancers.

KRAS Mutation Testing

Mutations in KRAS result in oncogenic constitutive activation of the RAS-RAF-MEK-MAPK signaling pathway (see Fig. 23.4). KRAS is one of the most frequently mutated oncogenes in human cancer, and mutations occur in 40% to 50% of colorectal cancers.²² Because KRAS sits downstream from EGFR, activation of KRAS has the potential to bypass the inhibitory effect of anti-EGFR therapy. Many studies have demonstrated that cetuximab is ineffective in patients whose colorectal cancers harbor KRAS mutations (with the possible exception of KRAS G13D).^64–66 KRAS mutation testing is a requirement of standardized chemotherapeutic protocols before cetuximab therapy is considered.^61,62

Technical approaches to KRAS mutation testing have been guided by the relatively narrow mutation spectrum in human neoplasms, with most recurrent mutations in colorectal cancer confined to codons 12 and 13 and less frequently to codons 61 and 146.²² In addition to PCR of the genomic region encompassing these codons followed by Sanger sequencing (Fig. 23.5), the limited profile of possible mutations allows for a variety of other mutation detection methods, especially mutation-specific PCR with panels of primers that correspond to all known recurrent mutations. Pathologists should be familiar with these methods primarily because some laboratories offer more comprehensive panels of mutation detection than others. Mutation analysis is readily performed on paraffin-embedded tissue, requiring only a small amount of tumor sample, and it is therefore applicable to biopsy samples. KRAS mutations are relatively homogeneous within colorectal cancers, most likely because mutations typically occur during the development of an adenoma and are therefore present in all subsequent generations of evolving malignant and metastatic clones.

The KRAS G13D mutation is one possible exception to the lack of response to anti-EGFR therapy in colorectal cancers harboring KRAS mutations. Several studies have shown that cetuximab therapy may have some efficacy in patients whose colorectal cancers harbor G13D mutations and that these mutations should not necessarily be considered a contraindication to anti-EGFR therapy.^67,68 Ongoing studies are assessing the magnitude of therapeutic benefit in this setting.

Similarly, many studies have looked at KRAS mutations only in codons 12 and 13, and there are few data regarding resistance in patients with mutations in codons 61 and 146. Although the latter two codons are not considered in some chemotherapeutic guidelines, there is evidence that they confer resistance, and it seems reasonable to assume that they will be included in future guidelines after additional validation data are available.⁶⁹

NRAS mutations occur in approximately 3% of colorectal cancers and are mutually exclusive of KRAS mutations.²² There is some evidence to suggest that NRAS mutations are associated with resistance to cetuximab therapy, but the data are limited. NRAS mutation testing is not recommended in routine settings, and the mutation is not considered a contraindication to anti-EGFR therapy.⁶⁹ With additional validation, however, it seems reasonable to expect that the finding of an NRAS mutation will be a contraindication to anti-EGFR therapy. NRAS mutations mostly occur at codon 61, rather than codons 12 and 13, and they are readily detected by PCR and sequencing or other mutation-specific PCR approaches.

Acquired Resistance to Anti-EGFR Therapy

As with many other targeted therapies used in cancer patients, despite a good initial response to anti-EGFR therapy, acquired resistance eventually develops.^66,80 Although the discovery of at least two mechanisms of acquired resistance represents an exciting advance in the development of tailored colorectal cancer therapy, the current level of understanding does not warrant routine testing of tissue samples from colorectal cancer patients with emerging resistance to anti-EGFR therapy.

KIT and PDGFRA Mutations

Derived from interstitial cells of Cajal, GISTs share almost universal expression of the tyrosine kinase receptor KIT. Activating KIT or platelet-derived growth factor receptor-α (PDGFRA) mutations occur in 80% to 85% of tumors and 5% to 10% of tumors respectively. They result in activation of the mitogen-activated protein kinase (MAPK) and phosphatidylinositide 3-kinase (PI3K) pathways. Overexpression of insulin-like growth factor receptor (IGFR) or direct mutation of BRAF occurs in some of the tumors with wild-type KIT and PDGFRA. Testing for KIT and PDGFRA mutations is typically performed by PCR and sequencing.

Imatinib Therapy

Imatinib mesylate (Gleevec, Novartis, East Hanover, N.J.) is an oral, small-molecule receptor tyrosine kinase inhibitor that works by blocking the adenosine triphosphate binding site of the constitutively activated mutant KIT or PDGFRA.⁸⁶ Binding effectively shuts down signaling. Since it was approved by the U.S. Food and Drug Administration (FDA) for use in GIST in 2002, imatinib has become first-line therapy for metastatic or unresectable GISTs. Approximately 80% of patients respond to initial imatinib therapy; however, the development of resistance is common, and progressive disease develops in more than one half within 2 years.

The type of mutation in KIT or PDGFRA is the best predictor of response to initial imatinib therapy (Table 23.5).⁸⁶ Approximately 60% to 80% of KIT mutations occur in the exon 11 intracellular juxtamembrane domain, and these mutations are associated with the best response to imatinib. Approximately 5% to 20% of mutations occur in the exon 9 extracellular domain, and they are associated with resistance to imatinib. Rare remaining mutations occur in the exon 13 ATP-binding pocket or exon 17 activation loop, and they also confer resistance to imatinib. In PDGFRA, more than 90% of mutations occur in exon 18, which is structurally analogous to KIT exon 17 and also resistant to imatinib.

Overall, approximately 10% to 15% of patients have primary resistance to imatinib therapy, defined as a lack of response or stable disease for less than 6 months. Most patients have imatinib-resistant mutations in exon 9 of KIT or exon 18 of PDGFRA or have no mutation.⁸⁶ Secondary resistance develops in patients whose tumors are initially sensitive to imatinib, typically at a median of 18 to 24 months. Secondary mutations in exons 13, 14, or 17 are found in approximately 40% to 80% of those developing secondary resistance.⁸⁶ Tumor heterogeneity is also seen, with different mutations identified in different lesions or within a single metastasis. Other mechanisms of secondary resistance include amplification of KIT and pharmacokinetic resistance.

Surgical resection or ablation is recommended in patients who are initially responsive but in whom limited progression occurs. By removing the imatinib-resistant clone, therapy may be continued in otherwise responsive disease. Dose escalation should be considered and may benefit as many as 30% of resistant patients.

Sunitinib Therapy

Sunitinib malate (Sutent, Pfizer Oncology, New York, N.Y.) is an oral, multitargeted receptor tyrosine kinase inhibitor that is active against KIT, PDGFRA, and other receptor tyrosine kinases. It was approved by the FDA in 2006 for use in patients with advanced GIST that is resistant to imatinib therapy and for those who did not tolerate imatinib.

Similar to imatinib, sunitinib binds the ATP-binding pocket and prevents tyrosine kinase activation. Sunitinib has a smaller molecular profile than imatinib, allowing greater activity against exon 13 and 14 KIT mutations and a broader range of exon 9 KIT mutations and wild-type KIT. Exon 17 KIT mutations and exon 18 PDGFRA mutations remain resistant. Overall, approximately 50% of patients with imatinib-resistant disease can benefit from sunitinib therapy, although the median progression-free survival is only 6 months.⁸⁶ Mutational analysis of imatinib-resistant disease has been recommended by some authorities to better select patients for sunitinib therapy. Currently, sunitinib is the only agent that is FDA approved for second-line therapy in GIST.

Other Targeted Therapies

Nilotinib has shown some activity against GIST, but because it does not demonstrate any clear benefit over imatinib, further development of this compound for use in GIST is unlikely. Sorafenib and regorafenib show promising results and have greater activity than imatinib against exon 9 KIT mutations and secondary KIT mutations in exons 14 and 17.⁸⁶ Regorafenib is effective in advanced GIST that has failed other standard therapies.⁸⁷ Many other receptor tyrosine kinase agents are being evaluated in a variety of clinical trials.⁸⁶ Targeting of downstream signaling has also been considered, including mTOR inhibitors, PI3K inhibitors, and anti-IGF1R therapies.

Resistance Monitoring during Therapy

Resistance to imatinib therapy eventually develops in most patients with GIST, even in those whose tumors are initially sensitive and have good response. Monitoring for the emergence of resistant KIT mutations plays an important role in therapeutic monitoring and decision making regarding second-line therapy. Unfortunately, the location and multiplicity of resistant disease makes biopsy sampling impractical. Emerging technologies that can detect KIT mutations in cell-free fluid samples such as serum and plasma offer significant gains in clinical utility in this setting. One such technology, BEAMing (see Rare Mutation Detection), was able to detect secondary resistant KIT mutations in the plasma of 48% of patients who had developed resistance to imatinib or sunitinib, compared with detection of secondary mutations in only 12% of tissue samples.¹⁴ This so-called liquid biopsy offers the potential for tailored monitoring of emerging resistance in any targeted therapy setting.

Light-Chain Restriction

The two types of light chains in humans are κ and λ. A single B cell expresses only one class of light chain, and it remains fixed for the life of the lymphocyte. Immunohistochemistry or in situ hybridization is used to determine the mixture of κ and λ light chains expressed in a non-neoplastic population of B cells. In contrast, a monoclonal population of B cells expresses one predominant light chain, a phenomenon known as light-chain restriction. Examination of light-chain expression is useful for determining whether a given B-cell population is reactive or neoplastic.

Southern Blot

Southern blot analysis is the traditional method for identifying clonal heavy-chain gene rearrangements, and it was one of the first routinely used tests in molecular diagnostics. DNA from normal tissue and lesional tissue is digested with restriction enzymes, electrophoresed on a gel, transferred to a membrane, and hybridized with a probe to the joining region of the IGH gene. Normal tissue and polyclonal lymphoid tissue reveal the normal banding pattern generated from the known restriction sites, whereas a clonal lymphoid population reveals an abnormal, rearranged monoclonal band. The major limitation of Southern blot analysis is that it requires 20 to 30 µg of high-quality DNA, which usually is not available unless specifically collected for this analysis.

Polymerase Chain Reaction

PCR is the method of choice for detecting gene rearrangements. Different approaches vary in their ability to detect many rearrangements or all possible rearrangements. The general approach is to combine one forward PCR primer with a variety of reverse primers corresponding to different regions that may be rearranged. In a polyclonal lymphoid population, there are innumerable rearrangements, yielding a cloud or smear when the PCR products are separated by electrophoresis. The presence of a dominant monoclonal population results in a distinct PCR band.