This method can be used to further increase sensitivity and specificity of a PCR. It is used frequently in hematology to detect a very small quantity of input molecules, in other words for minimal residual disease. Here, nested PCR is applied most often to determine fusion transcripts produced by a chromosomal rearrangement.
PCR testing can be conducted using any material. Between 5 and 10 ml of bone marrow with heparin, EDTA or citrate anticoagulant, i.e. peripheral blood, are needed for the analysis. Peripheral blood is sufficient as a test material, provided that malignant cells have passed into the blood. Therefore, bone marrow should be tested if passage has not occurred.
Nested PCR involves taking a small sample from a first PCR and using it as input material for a new PCR. Oligonucleotides that hybridize within the first amplification are used as primers. This additional amplification drastically increases sensitivity. In this case – depending on the type of mutation and the input material – it is possible to detect one malignant cell in 104 to 108 the number of healthy cells. Therefore, nested PCR is currently the most sensitive method to detect minimal residual disease for most target sequences. But this method also has the greatest risk of contamination, precisely because of its extreme sensitivity. It is essential to apply a variety of precautionary measures and always to conduct control reactions in order to avoid contamination.
Quantitative real-time PCR is another method for the amplification and detection of PCR products. Unlike the other detection methods, it is not a qualitative endpoint analysis, and is instead measured in real time during propagation of the PCR. Analysis of the logarithmic phase of this amplification permits precise quantification of the target sequences in the test material.
Real-time PCR testing can be conducted using any material. Between 5 and 10 ml of bone marrow with heparin, EDTA or citrate anticoagulant, i.e. peripheral blood, are needed for the analysis. Peripheral blood is sufficient as a test material, provided that malignant cells have passed into the blood. Therefore, bone marrow should be tested if passage has not occurred.
The method is based on introducing fluorescence-marked probes to the reaction mixture in addition to the specific primers required for PCR. During PCR, they hybridize with the continuously propagating amplification products and therefore emit fluorescence signals that are detected by the optical unit. Therefore, provided the specific target sequence is present, a rise in fluorescence intensity will occur directly during PCR. The point (PCR cycle) at which fluorescence above background levels is detectable for the first time correlates with the number of molecules that can be identified in the input material. The number of identified molecules of the target gene are benchmarked against a constant gene or transcript (normalized), which permits calculation of the number of malignant cells that the input material contains. Special PCR equipment fitted with optical units is necessary in order to conduct this detection method. While this method can also be used for the normal identification of a PCR product, real-time PCR reveals its true strength in particular during progress testing, e.g. during treatment, as it provides an opportunity for precise quantification and hence is used as a tool to measure treatment success. Real-time PCR also achieves extremely high sensitivities, so that – depending on the type of mutation and the input material – one malignant cell can be detected in 104 to 105 the number of healthy cells.
In many cases, targeted diagnostics will require the determination of the precise base sequences of a gene segment by means of sequence analysis.
Sequencing can be conducted using any material. Between 5 and 10 ml of bone marrow with heparin, EDTA or citrate anticoagulant, i.e. peripheral blood, are needed for the test. Peripheral blood is sufficient as a test material, provided that malignant cells have passed into the blood. Therefore, bone marrow should be tested if passage has not occurred.
Sanger sequencing involves the performance of sequencing reactions in which nucleotides are added to a PCR-based reaction. Each of the four different nucleotides carries a separate fluorescent marker and dideoxy group, which precipitate a break in the PCR chain reaction. The products are then placed in a matrix, e.g. a gel or a polymer, that separates them according to length, and they are then read by an optical unit to determine the precise base sequence in each case.
In a limited number of patients, Sanger sequencing can be implemented as the primary method for individual gene segments. The sensitivity of this method is 10–20% mutational load. Direct sequencing can be extremely laborious and expensive if several regions are selected for screening in a single patient, or if certain genes are screened for a large number of patients. There is a variety of screening methods available that permit the identification of whether mutation is present in certain gene segments, without determining the precise sequence (e.g. fragment length or melting curve analysis). Alternatively, mutation analysis can also be performed using next-generation sequencing, which permits a significantly larger throughput.
Hematological diagnostics faces significant challenges due to the rising diversity of clinically relevant molecular markers. Extremely sensitive, parallel analysis of hundreds of thousands of genome regions became possible with the introduction of modern, high-throughput sequencing methods, a.k.a. next-generation sequencing (NGS). This opens up new opportunities in mutation analysis and for progress control of hematopoietic neoplasias. For instance, NGS is used for BCR-AB1 mutation analysis among CML patients or to prepare comprehensive genetic profiles during diagnosis and prognosis of AML or MDS.
NGS testing can be conducted using any material. Between 5 and 10 ml of bone marrow with heparin, EDTA or citrate anticoagulant, i.e. peripheral blood, are needed for the test. Peripheral blood is sufficient as a test material, provided that malignant cells have passed into the blood. Therefore, bone marrow should be tested if passage has not occurred.
Next Generation Sequencing permits a staggering parallelization of the sequencing process. Methods based on amplicon deep sequencing or targeted enrichment are particularly suitable for routine diagnostics. Depending on the entity, the methods can sequence relevant gene segments with a high degree of sensitivity (1–3% mutational load). Panel testing permits the analysis of several hundred genes or genetic hotspots in one test cycle in less than a week. The findings are assessed using complex data processing and sequencing variants are then compared with SNP and mutation databases.
Several modern NGS platforms are available from routine diagnostics, for instance at Illumina or Thermo Fisher.
Array technology is a useful addition to classic cytogenetics. Array-based copy number analyses have become increasingly important in recent years. Application of this methodology permits whole genome, high-resolution detection of gains and losses in chromosomal material, including cytogenetically cryptic microdeletions and microduplications. It can also identify and characterize breakpoints and unknown fusion partner genes for unbalanced translocations. Screening for copy-neutral loss of heterozygosity (CN-LOH) is also possible. Frequently encountered synonyms for genomic array analyses include array CGH (array-based comparative genomic hybridization) and SNP array (single nucleotide polymorphism-based array).
Given that the detection of balanced translocations or intrachromosonal rearrangements is not possible without copy number changes, array technology is a useful addition to chromosome analysis and fluorescence in situ hybridization (FISH), although it cannot replace the other two techniques. Array technology is used as a complementary method, above all for patients with cytogenetically normal findings or extremely complex karyotype mutations.
Table 1: Advantages and limitations of genomic array analyses
Insight into gains and losses in the whole genome
Balanced rearrangements cannot be detected
Independent of malignant cell proliferation
Small clones (<15%) cannot be detected safely
Very high resolution
Copy-neutral LOH are detectable
Genomic array analysis involves the hybridization of patient DNA on an array. Additional hybridization of reference DNA may be necessary, irrespective of the technology used. Resolution of the genomic array varies depending on the particular array platform; the array we currently use provides resolution of approximately 100 kb. After around 48 hours of hybridization, the arrays are scanned and then characterized for mutations in the patient genome using a variety of bioinformatic algorithms.
In addition, the genomic array platform used in MLL research projects permits the identification of regions featuring copy neutral loss of heterozygosity (CN-LOH) for the validation of genomic gains and losses. Early studies have demonstrated that CN-LOH regions of tumor cells are frequently encountered in AML, myeloproliferative neoplasias and myelodysplastic syndromes and that they are associated with loss-of-function mutation for genes in the affected CN-LOH region. Neither chromosome analysis nor FISH have the capability to detect CN-LOH.