Molecular Diagnostics of Tubal Gut Neoplasms

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Chapter 23

Molecular Diagnostics of Tubal Gut Neoplasms

Mark Redston

John Iafrate

Introduction

Molecular diagnostic assays comprise three broad categories. Molecular microbiology is used for infectious disease diagnosis and classification. Genetic testing is applied to germline DNA for detecting heritable diseases, pathogenic mutations, and disease-associated single nucleotide polymorphisms. Molecular oncology methods are used for testing somatic tumor cell samples or their derived cellular or molecular constituents. This chapter focuses on the methods and applications of molecular oncology to the gastrointestinal (GI) tract and highlights the genetic and infectious disease testing used in the clinical practice of diagnostic pathology (Table 23.1).

Table 23.1

Validated Molecular Genetic Tests in GI Pathology

Disease State Test Method Clinical Application
Hereditary GI cancer Multiple genes IHC, MSI, PCR and sequencing Determine germline mutation status
Gastric and GEJ adenocarcinoma HER2 overexpression
HER2 amplification		 IHC
FISH, CISH		 Protein overexpression or gene amplification predicts response to anti-HER2 therapy
Colorectal adenocarcinoma Microsatellite instability testing PCR High-frequency MSI predicts possibility of Lynch syndrome, good prognosis, and lack of response to 5-FU
Mismatch repair protein expression IHC High-frequency MSI predicts possibility of Lynch syndrome, good prognosis, and lack of response to 5-FU
BRAF mutation analysis PCR ± sequencing BRAF mutation predicts MLH1 methylation in mismatch repair–deficient cancers and possible decreased response to anti-EGFR therapy
MLH1 promoter methylation Quantitative methylation-specific PCR Presence of MLH1 methylation excludes Lynch syndrome in MLH1-deficient cancers
KRAS mutation analysis PCR ± sequencing Absence of KRAS mutation predicts sensitivity to anti-EGFR therapy
Oncotype Dx colon cancer assay; ColoPrint Multimarker assay Can predict high risk of recurrence in stage II/III disease
GIST KIT mutation analysis PCR and sequencing KIT exon 11 mutations predict sensitivity to imatinib
PDGFRA mutation analysis PCR and sequencing PDGFRA mutations predict resistance to imatinib therapy
Lymphoma Translocation detection FISH, PCR Presence of specific translocations supports lymphoma diagnosis and classification
Infectious disease HPV, CMV, EBV CISH Diagnosis of specific infectious agent
Mycobacterium PCR Detection of organisms from formalin-fixed, paraffin-embedded tissue with negative AFB stains

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AFB, Acid-fast bacillus; CISH, chromogenic in situ hybridization; CMV, cytomegalovirus; EBV, Epstein-Barr virus; EGFR, epidermal growth factor receptor; FISH, fluorescence in situ hybridization; 5-FU, 5-flurouracil; GEJ, gastroesophageal junction; GI, gastrointestinal; GIST, gastrointestinal stromal tumor; HER2, human epidermal growth factor receptor 2; HPV, human papillomavirus; IHC, immunohistochemistry; KIT, CD117; MSI, microsatellite instability; PCR, polymerase chain reaction; PDGFRA, platelet-derived growth factor receptor-α.

Methodologies

Sample Collection and Processing

Most routine genetic testing is performed on high-molecular-weight genomic DNA and RNA extracted from white blood cells in whole blood samples. Improved methods use more limited samples, such as blood spots and buccal smears. Because a thorough discussion of all methods is beyond the scope of this chapter, we focus on testing of surgical pathology samples and other samples that may harbor neoplastic cells, DNA, or microsomes that may be used for molecular screening and diagnosis.

Tissue

Historically, extraction of high-quality DNA and RNA has required fresh or frozen tissue samples. If processed very quickly after surgical removal, these samples provide excellent nucleic acids for subsequent investigations. However, degradation of nucleic acids may occur. Degradation depends on several factors: tissue necrosis, duration of acute surgical ischemia, the time from resection to tissue harvesting, DNase and RNase activity during extraction, and improper storage. Attempts to use formalin-fixed, paraffin-embedded tissue samples create unique challenges and usually are less successful because of DNA fragmentation, DNA cross-linking, and contaminants. However, these samples are much more widely available, and improvements in nucleic acid recovery and subsequent testing have greatly increased the utility of these samples.

Microdissection

Tumor samples, particularly from carcinomas, are a complex mixture of neoplastic cells, stromal cells, and inflammatory cells. Some carcinomas, especially those of the pancreas and biliary tree, are characterized by scattered, small neoplastic glands embedded in an abundant desmoplastic stroma. Other cancers may be infiltrated by a dense lymphocytic response. As a result, unenriched tumor samples may vary in neoplastic cellularity from a high of approximately 80% to a low of only 10% to 20% tumor cells. The success of some molecular diagnostic assays depends on a neoplastic cellularity of at least 50%, and microdissection may be required to enrich the sample for neoplastic cells. Manual microdissection involves trimming frozen tissue blocks under the direct guidance of frozen section histology or scraping specified regions of neoplastic cells from an unstained section of paraffin-embedded tissue that is aligned with the adjacent hematoxylin and eosin (H&E) staining.1 These approaches can produce marked improvements in neoplastic cellularity.

Laser microdissection methods may be used to dramatically enrich neoplastic populations, although the yield of tissue is significantly reduced in these labor-intensive situations.1 Assay requirements must be tailored to the tissue parameters to determine which enrichment methods are needed. In liquid-phase enrichment, cells are digested to release them in a suspension and then purified by bead-tagged antibodies that target cell surface antigens. These approaches are common in research applications but are used much less frequently in clinical practice.

In addition to tissue samples obtained from sections cut from paraffin blocks, cytologic preparations may be used in molecular diagnostic assays. The number of cells and quantity of DNA available for testing are usually limited with these methods. The use of cytologic specimens in molecular assays needs to be closely supervised by a pathologist because the process may consume the permanent diagnostic record.

Circulating DNA

Small amounts of detectable, cell-free DNA circulate in the plasma and serum of healthy individuals and in persons with a variety of disease states. In patients with cancer, this includes DNA and RNA from tumor cells. Detection of a variety of tumor mutations, microsatellite alterations, methylation abnormalities, and chromosomal alterations has laid the foundation for development of assays for use in cancer screening, diagnosis, and therapeutic monitoring.2 In a similar manner, DNA from other biofluids (e.g., effusions, fecal samples, biliary aspirations, luminal washings) may be used in molecular diagnostic assays. These findings and technologies are an extremely active area of research and hold promise for the development of a variety of novel assays, but there currently are no validated tests in routine use for GI diseases that use these approaches.

Nucleic Acid Extraction and Purification

Traditional extraction of good-quality high-molecular-weight DNA requires a three-step extraction procedure. First, cell and tissue lysis use detergents, proteinase K, and RNase. Second, proteins are removed, avoiding RNA and chemical contamination. Third, DNA is purified. Conventional manual extraction techniques use phenol-chloroform purification and precipitation by using ethanol or isopropanol. Newer, kit-based approaches use a variety of bead-based elution technologies to purify DNA.3 Some methods yield DNA of lesser purity and lower molecular weight, although this result is acceptable for many applications. The methods used must appropriately balance ease of use, expense, quality, and downstream assay requirements.

RNA is extracted by two predominant methods: phenol-based extraction (e.g., TRIzol reagent) and silica matrix or glass fiber filter-based binding. TRIzol methods retain small RNAs, including microRNA (miRNA) and small interfering RNA (siRNA). TRIzol reagent also includes guanidine isothiocyanate to maintain the integrity of RNA while disrupting cells and dissolving cell components.

After extraction, nucleic acids must be accurately quantified to assess the success of the extraction and to determine the correct quantity of template for subsequent applications. Quantification is typically performed by using an ultraviolet spectrophotometer with a known control, usually calf thymus DNA.

Detection of Sequence Variants

Detection of single nucleotide changes and small insertions and deletions in an unknown nucleic acid sample is the most common assay performed in molecular diagnostics. These assays are performed to identify hereditary germline disease-associated mutations, single nucleotide sequence variants (i.e., polymorphisms), and somatic aberrations in tumor cells. Although sequencing is the gold standard method, it has not always been the most practical approach because of the size of the target gene that needs to be analyzed, the limited DNA sample that may be available, or the rarity of the variant sequence in the nucleic acid population to be analyzed. Many different approaches to screening for mutations have evolved in the past 20 years in response to improved technologies.

For simplicity, sequence variant detection methods may be divided into polymerase chain reaction (PCR)–based technologies and sequencing. Methodologic approaches may be tailored to the expected result. For instance, the approaches to detect a common known sequence variant at a set nucleotide (e.g., BRAF V600E mutation in colorectal cancer) are very different from mutation screens applied to an entire exon, gene, or panel of genes. PCR-based approaches use a variety of different techniques. Common approaches include PCR primers that are designed for specific recurrent mutations and real-time PCR melting curve analysis. As sequencing technologies have improved and reactions have become simpler and cheaper to perform and analyze, sequencing approaches have become the first choice for sequence variant detection.

Sanger Sequencing

Now known as first-generation sequencing, automated Sanger sequencing was introduced in 1977. It uses electrophoretic separation of randomly terminated linear sequence extensions and remains the most widely available technology today.4 It is accurate, has well-defined chemistry, and is best suited for reading 500-base pair (BP) to 1-kilobase (kb) DNA fragments in a single reaction. Modern platforms use automation, fluorescent chemistry, and capillary electrophoresis. They have served as the primary machine for single-gene diagnostics in molecular diagnostics laboratories and as the workhorse for the first generation of the genome sequencing project.

Sanger sequencing does have technical limitations, particularly throughput (i.e., number of bases per second that can be read). Electrophoretic separation is the predominant rate-limiting step, severely limiting the speed and cost of assays. For example, it has been estimated that one automated sequencer would require several decades and tens of millions of dollars to sequence a single genome.4

Next-Generation Sequencing

Also known as second-generation sequencing and massively parallel sequencing, next-generation sequencing (NGS) technologies are capable of sequencing large numbers of different DNA sequences in parallel (i.e., in a single reaction).4 NGS technologies were developed in part from focused investment by the National Human Genome Research Institute (NHGRI) in an effort to markedly reduce the expense of large-scale sequencing and thereby rapidly advance the realization of personalized medicine.

NGS assays the addition of nucleotides to immobilized and spatially arrayed DNA templates. Although the various platforms use four basic steps (i.e., sample collection, template generation, sequencing reaction, and detection), they use substantially different methods of template generation and sequence interrogation (Table 23.2). Because NGS represents the most important technologic development emerging in molecular diagnostics, it is considered in more detail in the following sections.

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.

Targeted Sequencing

NGS is used to target panels of genes that have relevant mutations in the diagnosis or management of disease. NGS becomes cost-effective compared with Sanger sequencing diagnostics when multiple genes need to be sequenced. Examples include panels of genes for the diagnosis of genetic diseases and for the diagnosis of oncogenic driver mutations in cancer.

Multigene oncogenic assays have been established for lung cancer, for metastatic disease that has failed other therapeutic options, and for a variety of research studies, and they have offered significant advantages to traditional molecular diagnostic approaches.6,7 A panel developed for screening known genes associated with colorectal polyposis and Lynch syndrome has been a powerful and cost-effective approach that eliminates the traditional stepwise approach to genetic characterization.8,9 GI tract, liver, and pancreaticobiliary malignancies do not have other sufficiently validated gene targets to warrant additional NGS gene panels, and although current applications are mostly limited to the research setting, this situation is expected to change dramatically in the next 5 years.

Whole-Exome Sequencing

Exome sequencing refers to large-scale sequencing of genomic coding regions, which comprise only approximately 2% of the genome. Exome sequencing may be directed at the whole exome or may target the approximate 3000 genes or smaller subsets known to be involved in human disease. Although exome sequencing has not been validated for routine clinical use, it has revolutionized research into the genetic basis of human disease and the comprehensive identification of novel gene mutations in human tumors.

Whole-Genome Sequencing

Although whole-genome sequencing (WGS) does not have routine validated clinical applications, it is technologically feasible on an individual basis and has been used with great success in selected cases and research studies. WGS has been used in the analysis of tumors and has resulted in the rapid identification of novel translocation fusion proteins not identified by other methods. For example, WGS has been used in a clinical trial setting for an individual patient with metastatic colorectal cancer in whom a novel experimental therapy was used based on the mutation profile of the tumor.10

Rare Variant Detection

In most studies and applications, NGS has been used to broadly sequence the entire genome or large portions of the genome to detect variants that are present at high frequency in the sample DNA. NGS also may be used to repeatedly sequence large numbers of the same template and identify rare variants. These deep sequencing studies use very dense coverage of a small portion of the genome. Examples of potential applications include identification of circulating donor DNA in patients with very early transplant rejection and identification of fetal Down syndrome in circulating DNA of the pregnant mother of the fetus.11 The technique has many potential applications in screening and early detection of cancer.

Rare Mutation Detection

Technologies designed to identify rare mutation loads in clinical biospecimens can be applied to screening for malignancy (e.g., detection of KRAS mutations indicating colorectal adenomas and carcinomas in fecal samples), monitoring of the cancer burden and minimal residual disease (e.g., reverse transcriptase PCR detection of leukemia-specific translocations in bone marrow and blood samples), and detection of secondary oncogenic mutations leading to resistance to targeted therapy (e.g., Beads, Emulsion, Amplification, and Magnetics [BEAMing] technology to identify resistance mutations in KIT). The potential superiority of these approaches in blood and plasma samples compared with tissue biopsies has generated a concept referred to as liquid biopsy.

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.

Detection of Translocations and Genomic Copy-Number Alterations

Chromosomal structural variations are a hallmark of cancer and include abnormal chromosomal number (i.e., aneuploidy), translocations, focal amplifications, and subchromosomal deletions.16 Although deletions and abnormalities in chromosome number are among the most common changes, particularly in solid tumors, translocations and focal amplifications have garnered the most interest because of their association with oncogenic activations that may be susceptible to targeted inhibition. Translocations and other rearrangements also are important in the routine characterization and diagnosis of hematologic malignancies.

Methods used in the characterization of chromosomal structural variations include traditional cytogenetics, fluorescence in situ hybridization (FISH), comparative genomic hybridization (CGH) array-based methods (e.g., CGH array), and PCR. The remainder of this discussion focuses on the methods most commonly applied to tissue samples in clinical molecular diagnostics: FISH and PCR. CGH arrays are briefly discussed (see Microarray-Based Platforms).

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.

Microarray-Based Platforms

Array-based hybridization is a technology that uses a large number of targets that are densely arrayed on a small, readable substrate, allowing multiple, simultaneous hybridizations. Array targets are immobilized on glass slides or other materials and may consist of DNA, cDNA, PCR products, oligonucleotides, RNA, or proteins.

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.

Multimarker Assays

The preceding discussions on sequencing and array technologies provide insight into the depth of genomic and gene expression data that can be harvested from analyses of a pathology specimen. For example, the Cancer Genome Anatomy Project recently concluded many years of work detailing the genetic and epigenetic aberrations of a panel of colorectal cancer tumors assayed by many technologic platforms.21 In combination with the efforts of the pharmaceutical industry to develop targeted therapies, research in the development of multimarker assays is likely to yield significant progress in the coming years.

Molecular Diagnostics of Hereditary Gastrointestinal Cancer

Molecular diagnostics were applied to hereditary GI cancer syndromes, with germline testing for familial adenomatous polyposis and Lynch syndrome beginning in the early and mid-1990s, respectively.22 In the past 20 years, hereditary cancer testing has expanded to include all of the polyposis syndromes and familial gastric cancer (Table 23.3).

Table 23.3

Molecular Genetic Diagnosis of Polyposis and Other Hereditary GI Cancer Syndromes

Entity Gene Proportion of Cases Attributable to Gene
Polyposis Syndromes
Familial adenomatous polyposis (FAP) APC >95%
Attenuated adenomatous polyposis (AFAP) APC 20-30%*
MUTYH (MYH)† 20-50%
Peutz-Jeghers syndrome (PJS) STK11 (LKB1) Familial 100%; simplex 90%
Juvenile polyposis (JPS) SMAD4 20%
BMPR1A 20%
Cowden syndrome PTEN‡ 85%
Bannayan-Riley-Ruvalcalba syndrome (BRRS) PTEN‡ 65%
Proteus or proteus-like syndrome PTEN‡ 20-50%
Nonpolyposis Hereditary Cancer
Lynch syndrome MLH1 50%
MSH2 40%
MSH6 7-10%
PMS2 5%
EPCAM (TACSTD1)§ 1-3%
Hereditary diffuse gastric cancer CDH1 30-50%
Genetic Basis Not Defined
Serrated polyposis syndrome Unknown
Familial colorectal cancer type X Unknown
Familial esophageal cancer Unknown

* The proportion of cases with APC mutations increases with increasing numbers of adenomas.

† AFAP caused by MUTYH mutations is also known as MUTYH-associated polyposis (MAP). Some MUTYH mutation carriers have more than 100 polyps at presentation, and these cases are differentiated from FAP by the absence of an autosomal dominant family history and patient age older than 35 years.

‡ The term PTEN hamartoma tumor syndrome (PHTS) is used to describe cases with PTEN mutations.

§ Hereditary EPCAM mutations cause Lynch syndrome by inducing MSH2 methylation and gene silencing in colonic epithelial cells.

With the exception of Lynch syndrome (discussed later), hereditary GI cancer molecular testing does not involve anatomic pathology specimens. Germline testing for hereditary cancer mutations is performed on genomic DNA extracted from white cells obtained from peripheral blood. Mutational testing is typically performed by PCR and Sanger sequencing or pyrosequencing. With the advent of other NGS technologies, much of this single-gene mutational testing is being combined into panels of genes, allowing significant reductions in cost and turnaround time.8,9

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