Genetics diagnosis

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Genetics diagnosis
2013-10-12 16:08:12

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  1. When do we perform prenatal genetic testing?
    • 1. A mother of advanced age (>35 years) because of greater risk of trisomies  
    • 2. A parent who is a carrier of a balanced reciprocal translocation, robertsonian translocation, or inversion (in these cases the gametes may be unbalanced, and hence the progeny would be at risk for chromosomal disorders)
    • 3. A parent with a previous child with a chromosomal abnormality  
    • 4. A fetus with ultrasound-detected abnormalities  
    • 5. A parent who is a carrier of an X-linked genetic disorder (to determine fetal sex)  
    • 6. Abnormal levels of AFP, βHCG, and estriol performed as the triple test.
  2. When do we perform postnatal genetic testing?
    • Multiple congenital anomalies  
    • •   Unexplained mental retardation and/or developmental delay  
    • •   Suspected aneuploidy (e.g., features of Down syndrome)  
    • •   Suspected unbalanced autosome (e.g., Prader-Willi syndrome)  
    • •   Suspected sex chromosomal abnormality (e.g., Turner syndrome)  
    • •   Suspected fragile-X syndrome  
    • •   Infertility (to rule out sex chromosomal abnormality)  
    • •   Multiple spontaneous abortions (to rule out the parents as carriers of balanced translocation; both partners should be evaluated)
  3. How does gene chips (microarray) work?
    Short sequences of DNA (oligonucleotides) that are complementary to the wild-type sequence and to known mutations are “tiled” adjacent to each other on the gene chip, and the DNA sample to be tested is hybridized to the array. Before hybridization the sample is labeled with fluorescent dyes. The hybridization (and consequently, the fluorescent signal emitted) will be strongest at the oligonucleotide that is complementary to wild-type sequence if no mutations are present, while the presence of a mutation will cause hybridization to occur at the complementary mutant oligonucleotide
  4. What are the three general categories of genetic testing?
    • Mutation detection of known sequence changes can be performed. This type of testing is targeted and typically limited to a predefined number of sequence changes, selected in advance. Selection is generally based on association with clinical phenotypes. Sequence changes may be located within a single gene or across multiple genes. Depending on the testing method used, the number of included sequence changes can range from a single mutation to thousands of mutations.
    • Cytogenetic studies of large structural variants are typically performed when the phenotype does not seem limited to point mutations and relatively small deletions and duplications. Such studies are helpful in syndromic phenotypes and for constellations of symptoms typically associated with abnormalities on the scale of chromosomes rather than single exons or genes.
    • Genotyping methods can be applied to identify mutations not selected in advance. These methods aim to discover mutations, and can target a gene with a known heterogeneous distribution of mutations, or can target larger segments of the genome to identify known or novel variations
  5. What are the methods for detecting known mutation?
    PCR, Amplification refractory mutation system, Allele specific oligonucleotide hybridization, Genotyping microarrays
  6. What are the methods for detection of cytogenetic abnormalities?
    Chromosomal analysis, FISH (Interphase FISH, spectral karyotyping), Array comparative genomic hybridization.
  7. What are the methods for GENOTYPING NEW MUTATIONS?
    Heteroduplex analysis, Single strand conformation analysis, Automated sequencing, Whole genome sequencing, Southern and Northern blotting.
  8. What are two technical components underlie the polymerase chain reaction (PCR) process?
    the existence of thermostable DNA polymerases purified or cloned from microorganisms living in hot springs and the ability to readily synthesize specific oligonucleotide primers of 20 to 30 residues.
  9. What does the standard PCR reaction mixture consists of ?
    • 1. a large excess of oligonucleotide primer pairs,
    • 2. a template DNA (typically genomic DNA), 3. free deoxynucleotide triphosphates (DNA bases),
    • 4. reaction buffer
    • 5. thermostable DNA polymerase (the enzyme that drives the PCR reaction).
  10. What are the stages of PCR?
    • The PCR consists of three stages that are repeated 30 to 40 times
    • 1. Denaturing — The mixture is heated to 95°C to allow the double-stranded template DNA to denature into single strands.
    • 2. Annealing — The mixture is cooled to a temperature just below the predicted primer pair melting temperatures, resulting in primer binding to the single-stranded DNA templates, followed by binding of DNA polymerase to the 3’ end of the primers.
    • 3. Elongation — The temperature is then raised to a polymerase activation temperature (~70 to 72°C) to initiate chain elongation. The polymerase catalyzes the addition of free nucleotides to the 3’ ends of the primers. Bases are added to complement the template sequence (A with T, C with G). Elongation continues for about one minute and is terminated by cycling back to step one of the reaction: heating the temperature to 95°C, resulting in strand separation.
  11. If the reaction starts with two copies, what is the yield after N cycles of PCR?
  12. How can highly-specific amplification be achieved In PCR?
    Highly-specific amplification can be achieved if the sequence combination of the primer pairs is present only once in the genome
  13. What is done after the PCR reaction is complete?
    • After the reaction is completed, the amplified product (called an amplicon) can be visualized by gel electrophoresis, during which the negatively charged DNA fragments migrate toward the positive electrode through a prepared gel.
    • The pores of the gel depend upon its agarose percentage and can be compared to a (molecular) sieve.

    • Thus, migration speed of the DNA fragments through the gel depends on both DNA fragment size and the applied voltage.
    • After electrophoresis and addition of an ethidium bromide stain, which forms a complex with nucleic acids, exposure of the gel to ultraviolet light demonstrates the amplified fragment as a single fluorescing band.
  14. Migration speed of the DNA fragments through the gel depends on ..................
    both DNA fragment size and the applied voltage.
  15. What are the advantages of PCR?
    • Rapidity — The amplification reaction takes approximately three hours. Additional time (several hours) is required to first extract DNA from cells and then perform gel electrophoresis after completion of PCR.
    • Extreme sensitivity — PCR can amplify the DNA from a single cell and can be used in preimplantation genetic diagnosis in embryos.
    • Robust — Degraded DNA can frequently be successfully amplified.
    • Specificity — The technique also permits the detection of small nucleotide mutations (see below), as well as trinucleotide repeat expansions.
    • Inexpensive and simple
  16. What are the disadvantages of PCR?
    • Because of its extreme sensitivity, contamination of the sample with small amounts of extraneous DNA may result in a spurious or false-positive finding. To help decrease the incidence of this problem, diagnostic PCR laboratories are often subdivided into separate designated pre- and post-amplification areas in different rooms.
    • To design the appropriate PCR primers, the nucleotide sequence information of the region of interest must be known.
    • The ideal length of a sequence to be amplified (PCR amplicon) is several hundred nucleotides. Although longer length PCR methods are available, the analysis of very large regions is more difficult and requires different techniques.
    • Similarly, large mutations, such as duplications or deletions that encompass the amplified region, cannot be detected because only the unaffected allele will be amplified
  17. What is the probability of a primer with N nucleotide to occur in human genome?
    • (3 X 109) x 1/4n  
    • where 1/4n is the probability of a given sequence of length N bases and the constant 3 X 109 is the number of bases in the human genome.Thus, a PCR primer 20 nucleotides in length would be estimated to occur by chance at a frequency of 0.003 per genome, and a primer 25 nucleotides in length would occur at a frequency of 0.000003 per genome. Thus, these estimates demonstrate that a specific 20-residue sequence is likely to be unique in the human genome
  18. Why is the specificity of PCR is very high?
    • A specific 20-residue sequence is likely to be unique in the human genome.
    • Furthermore, PCR requires two primers to colocalize with a specific orientation to one another in a relatively small genomic region. Therefore, the probability of amplifying more than one region is exceedingly small
  19. What is whole genome amplification?
    Whole genome amplification (WGA) non-specifically amplifies the entire genome and is used when sample quantities are too small to meet the desired application's requirements. Random, very short primers are used, along with Phi29 polymerase, a highly-active polymerase capable of incorporating over 70,000 nucleotides per priming event. In order to limit amplification bias (uneven amplification of different genomic sequences or alleles), it is best to amplify the template DNA only a few 1000-fold.
  20. What is the difference between PCR and WGA?
    WGA is distinct from standard PCR in that it is non-specific (all genomic sequences are targeted, not only one site), and it generates fragments that are tens of thousands of bases in length, compared to 500 to 1000 bases that are typical for PCR
  21. What is the precursor to most gene expression microarray experiments?
    Linear amplification of RNA
  22. Why linear amplification of RNA is preferred over exponential amplification (PCR) in gene expression microarray experiments?
    because occasional imbalances in sequence amplification rates (that result from preferential reaction kinetics and favor amplification of some regions over others) can bias subsequent quantification
  23. What is the general method for global amplification of mRNA?
    • It is based upon the Eberwine protocol, whereby single-stranded RNA is the template, the reaction is primed by a poly-T oligonucleotide primer that binds the 3’ poly-A tail common to most mRNA sequences, and the reaction is catalyzed by a reverse transcriptase which generates complementary DNA strands.
    • Following second strand synthesis with DNA polymerase, and subsequent sample purification, T7 polymerase is used to drive in vitro transcription (IVT), producing multiple copies of amplified RNA (aRNA) from each dsDNA. By incorporating labeled nucleotides in the in vitro transcription step, amplification products are also prepared to be used as probes in expression analysis. Amplification achieved is typically in the range of 10- to 100-fold, but can vary depending on the amount of input RNA
  24. What are the major clinical applications of PCR?
    genotyping and sequencing for diagnostic and predictive testing
  25. What is genotyping?
    Genotyping is the process of characterizing an individual’s genotype status (the combination of alleles) at a particular genomic location. Most types of genetic variation can be characterized using PCR techniques, including microsatellite and single nucleotide polymorphism (SNP) markers, insertion/deletionvariants (indels), and some structural variants such as copy-number variants
  26. What is the base technique for SNP genotyping?
  27. What is DNA sequencing?
    DNA sequencing is the process of characterizing the nucleotide (base-pair) sequence of a specific DNA sequence. Clinically, sequencing is used to identify the pathogenic mutations in individuals with specific genetic disorders caused by rare variants at a specific locus. An example is resequencing of the BRCA1 locus in individuals at risk for familial breast cancer. Current clinical sequencing is PCR-based, with each assay focused on a specific gene region.
  28. What method is used for identifying rare sequences such as searching for DNA rearrangements in the setting of neoplasia or prenatal diagnosis?
  29. What is non-invasive prenatal diagnosis (NIPD)?
    NIPD relies on detection of free fetal DNA (ffDNA) in maternal blood;
  30. What is Quantitative PCR?
    Real-time quantitative PCR (qPCR) is a highly-sensitive method for quantifying the absolute or relative amount of a specific nucleic acid sequence in which the accumulation of PCR products over time is measured directly, without post-PCR modifications
  31. What is the application of quantitative PCR?
    gene expression profiling, quantification of viral load, and copy number variation genotyping
  32. How is qPCR performed?
    • qPCR is performed by quantifying the amount of amplified product with each round of PCR cycling. In addition to the two primers that are necessary for successful PCR amplification, this method applies a fluorescent, non-extendible probe that hybridizes to the target sequence between the primer pair.
    • This probe contains a fluorogenic reporter dye at its 5' end and a quencher at the 3' end. The quencher blocks fluorescent emission as long as it is in close proximity to the reporter. As the primer extends downstream during amplification, however, the exonuclease activity of the DNA polymerase cleaves away the hybridized probe and removes the quencher. An increase in fluorescent signal ensue
  33. How is QPCR performed?
    • The mechanism of quantitative real-time PCR, which is based upon a probe that contains a fluorogenic reporter dye at its 5' end and a quencher at the 3' end (shown as a black line with a white box and a Q at its 5' and 3' end, respectively). With this technique, the tagged probe, which is designed to hybridize to one strand of the target DNA sequence (blue line), is added to a normal PCR. The quencher blocks fluorescence emission as long as it is in close proximity to the reporter. As the primer extends downstream during amplification (extended green arrow), the exonuclease activity of the DNA polymerase cleaves away the hybridized probe and removes the connection between the fluorescent dye (black box) and the quencher (Q). An increase in fluorescence signal ensues as more amplicons are generated.
  34. How is qPCR curve interpreted ?
    Due to the exponential nature of the PCR, the fluorescent signal increases proportionally to the amount of generated PCR product until a plateau is reached. Quantitation is accomplished by comparing the cycle number at which the patient sample reaches a predetermined level of fluorescence to a standardized curve of a control sample, thus deriving copy number at the start of the reaction. This method is rapidly becoming the method of choice for monitoring residual neoplastic disease in patients receiving chemotherapy and/or hematopoietic cell transplantation
  35. qPCR interpretation
    • The polymerase chain reaction (PCR) can be quantitated in its early exponential phase. Panel A: If there are many targets in the starting sample (eg, curves on the left), the generated fluorescent signal will cross a predetermined threshold value (CT) early in the PCR cycling process. If there are few targets (eg, curves on the right), the signal will rise more slowly and will cross CT at a later PCR cycle. Panel B: When CT is plotted versus the starting quantity of target DNA, the relationship is linear over more than 8 logarithms. To establish the amount of targets in a sample, one can simply determine the cycle number at which CTis crossed and use the standard curve to infer the number of targets present in the sample. Panel C: In an example of a cancer patient with t(14;18), the initially large number of targets present (red curve) is markedly reduced after treatment (blue curve).
  36. What are the advantages of qPCR?
    Extreme sensitivity and a wide dynamic rangeTime-efficient, as the method requires no post-PCR processing of samples
  37. How can gene expression profiling be done with qPCR?
    • Specific gene transcripts can be measured by qPCR from RNA samples that have been converted to cDNA by reverse transcription. Typically, relative quantification is used to calculate fold-change among a set of samples
    • For example, inflammation-related genes were assayed by Taqman qPCR, comparing normal tissue with colon cancer samples
    • qPCR expression profiling is used to validate measurements of specific candidate genes identified through genome-wide expression microarray studies
  38. Viral load can be measured using.......
  39. How can targeted CNV genotyping be done?
    • Copy number variants (CNVs) are large (>1 kb) genomic regions showing differential copy number among individuals .
    • Real-time qPCR is a robust and relatively high-throughput method for targeted CNV genotyping
  40. Which method is often used for monitoring residual neoplastic disease in patients receiving chemotherapy and/or hematopoietic cell transplantation?
  41. How can Restriction enzyme digestion in PCR be used for diagnosis?
    • Restriction enzyme digestion can be used to detect mutations that create or destroy a restriction enzyme site. Specific restriction enzymes, usually isolated from bacteria, recognize unique short sequences within a DNA fragment. These enzymes can cleave the DNA strands at that exact site. If a mutation either changes the DNA code to a sequence that creates a new restriction enzyme site or obliterates an existing restriction enzyme site, the mutation can be detected based upon the presence or absence of the site
  42. What are the advantages and disadvantages of restriction site diagnosis?
    • Advantagestechnically easy, Restriction enzyme analysis detects specific mutations and can be applied to many samples concurrently.
    • Disadvantages: impractical for disorders caused by a large number of different mutations and for the detection of mutations associated with nucleotide sequences that require the use of expensive restriction enzymes, Only a small fraction of existing point mutations actually create or remove a restriction site.Incomplete digestion can produce erroneous results.
  43. Which methods other than PCR can be used to detect known mutation?
    • Amplification refractory mutation system
    • Allele specific oligonucleotide hybridization
    • Genotyping microarrays
  44. What is Amplification refractory mutation system?
    • can be used to detect known point mutations
    • requires a multiplex PCR reaction
    • two primer pairs are added to a single PCR tube and two separate sequences from one piece of DNA are amplified in the same reaction
    • One reaction (using the control primer pair) is an internal control to demonstrate that the PCR reaction itself has worked. The other reaction (using a primer pair specific for the mutation under study) will amplify the target sequence depending upon the presence or absence of a specific point mutation
    •  A second tube contains DNA from the same patient, and includes the control primer pair and a primer pair that amplifies only the normal sequence. The only difference between the primers for the normal and mutant sequence is complementarity of one primer at the 3' end, where one is identical to the normal and one to the variant sequence. The other primer is the same for both reactions, and the product size will be the same.
    • When run side by side on a gel, these samples will exhibit homozygosity for the normal sequence, homozygosity for the mutation, or heterozygosity
  45. ARMS example
    • This example of amplification refractory mutation system (ARMS) PCR shows how this technique can be used to distinguish between an individual who is homozygous for a normal gene and individuals homozygous or heterozygous for mutation in that gene. In this case separate primers are designed to match the normal gene sequence or the mutant gene sequence. The DNA sequences in red represent the patient DNA; the sequences in green are the primers. The box shows the mutant base and the corresponding altered base on the mutant primer.
    • A an agarose gel is run with the PCR products, and the presence of a band shows whether the normal or mutant DNA sequence was present.
    • Lanes 2 and 3: Individual with the normal gene; only the normal primer gives a PCR product.
    • Lanes 4 and 5: Individual homozygous for the mutation; only the mutant primer gives a PCR product.
    • Lanes 6 and 7: Individual heterozygous for the mutation; both the normal and mutant primers give PCR products.
    • Note: A control DNA product was present in all lanes, verifying that the PCR reaction worked.
  46. What are the advantages and disadvantages of ARMS?
    • Advantages: Easy, can be used to assess specific point mutations and assess many samples concurrently
    • Disadvantages: impractical for disorders caused by a large number of mutations, Primer pairs must be designed for all reactions, every patient sample require multiple PCR tubes
  47. What is Allele specific oligonucleotide hybridization?
    • Allele-specific oligonucleotide (ASO) hybridization involves the placing ("spotting") of denatured PCR-amplified DNA onto a membrane and subsequent hybridization with short allele-specific, labeled probes.
    • Under optimal hybridization and washing conditions, hybridization will only occur if the probe sequence is perfectly complementary to the single-stranded sample DNA
  48. How does ASO work?
    • Typically, PCR products from one patient sample are fixed onto two identical membranes (a "dot-blot"), one of which is hybridized with a probe that contains the normal sequence, while the other is hybridized with a probe for the mutant sequence.
    • The two probes should differ by just one nucleotide, corresponding to the point mutation under investigation.
    • After exposure to an autoradiographic film in the case of radioactive probe labeling, or after chemical treatment in the case of biotinylated oligomers, positive signals are scored and heterozygosity or homozygosity for the normal or mutant sequence can be determined
  49. ASO
    • This representation of ASO hybridization with dot blot analysis shows how this technique can be used to distinguish between an individual who is homozygous for a normal gene from individuals homozygous or heterozygous for a mutation in this gene. Patient DNA is red, and probes are green. In this case a normal gene is detected by a probe with the normal sequence, and a mutant gene is detected by a probe with a mutant sequence. The point mutation is denoted by an "X".
    • Box 1: the normal probe is hybridized to spots of DNA "dotted" from a normal individual, an individual homozygous for the mutation, and an individual heterozygous for the mutation. The probe gives a signal for the normal and heterozygous individuals, but not the homozygous individual (who lacks the normal gene).
    • Box 2: the mutant probe is hybridized to spots of DNA "dotted" from a normal individual, an individual homozygous for the mutation, and an individual heterozygous for the mutation. The probe gives a signal for the homozygous and heterozygous individuals but not the normal individual (who lacks the mutant gene).ASO: allele specific oligonucleotide.
  50. What are the advantages and disadvantages of ASO hybridization?
    • Advantages: suitable for analysis of specific mutations or polymorphisms in numerous samples, highly sensitive and specific 
    • Disadvantages include: Each ASO probe can only detect one specific sequence, ASO hybridization is amenable to small DNA mutations only.
  51. What is genotyping microarray?
    • They can interrogate a flexible number of mutations at the same time in one or multiple genes, for one or many different patients at the same time.
    • Automated
    • Advantages: suitable for high-throughput analysis of specific mutations or polymorphisms in numerous samples, Relatively less hands-on work is required per sample.
    • Disadvantages :  expensive, not suitable for low-volume testing, in particular when the microarrays can be used only one time, regardless of whether only one or many samples are tested
  52. What are cytogenetic abnormalities and how can they be detected?
    • Cytogenetic abnormalities are genetic defects that involve large regions of chromosomes rather than small pieces of DNA (ie translocations, large deletions, or aneuploidies).
    • These defects can be detected by at least three methods – chromosomal (karyotypic) analysis, fluorescence in situ hybridization (FISH) using specific DNA probes on either metaphase chromosomes or interphase nuclei, and array comparative genomic hybridization (aCGH). 
  53. Chromosomal analysis (also called chromosome banding) is used to detect .........................................
    changes in large regions of chromosomes (translocations, large deletions, or aneuploidies)
  54. How is chromosomal analysis performed?
    • To perform chromosome analysis, lymphocytes, usually obtained from the peripheral blood, are cultured in vitro and stimulated to divide under the influence of mitogens.
    • Other cell types, such as amniocytes, bone marrow cells, fibroblasts, and tumor cells can also be analyzed, often without a mitogenic stimulus.
    • Once the cells divide readily, a chemical is added to arrest mitotic division in metaphase. In this phase of the cell cycle, the chromosomes are maximally contracted and hence their banding patterns are easier to recognize
    • Chromosomal banding is then used to identify each individual chromosome, for assessment of whether the correct number of each chromosome is present (two of each autosome plus the sex chromosomes) and whether there are structural abnormalities.
    • This technique can be performed using various enzymes and dyes. The most frequently used banding technique is GTG (G-banding with trypsin and Giemsa-banding).
    • An identical banding pattern is seen in Q-banding, in which the chromosomes are stained with a fluorescent dye and viewed under ultraviolet illumination
  55. Where and why is chromosomal banding used?
    • Used in chromosomal analysis
    • used to identify each individual chromosome, for assessment of whether the correct number of each chromosome is present (two of each autosome plus the sex chromosomes) and whether there are structural abnormalities
  56. The typical resolution of chromosome banding is about ........... in a haploid set of 23 chromosomes
    400 bands
  57. What is high resolution banding?
    If higher banding resolution is desired to study relatively small chromosomal rearrangements, the cell cycles can be synchronized and cells arrested in prometaphase (550 bands) or even prophase (approximately 800 bands)
  58. What are the advantages and disadvantages of chromosomal banding?
    • Advantages :In contrast to molecular genetic studies, chromosome banding techniques show the entire genome at one time/  suitable in diagnostic situations where a specific anomaly is suspected (eg, the Philadelphia chromosome in chronic myeloid leukemia, CML)/ In disorders having deletions of varying size within a specific chromosome, such as multiple myeloma, karyotypes from many patients can be compared, in order to find the critical disease-associated region
    • Disadvantages are: Most chromosome banding techniques can only detect major structural abnormalities and will not detect smaller regions of DNA gain or loss/ Interpretation is labor intensive and highly dependent upon operator experience and skill
  59. Chromosomal 13 del in MM
    • Conventional cytogenetic analysis with G banding was performed in 106 patients with multiple myeloma. Each vertical line to the right of the figure (the long arm of chromosome 13) indicates the part of chromosome 13 that was found to be deleted in one patient. With the exception of eight patients, the minimal region of deletion overlap appears to be in the 13q14 region (red arrow).
  60. FISH can be done in which phases of cell cycle?
    metaphase chromosomes or interphase nuclei
  61. Metaphase FISH can be used for.........
    identification of large chromosomal abnormalities, including deletions, duplications, and translocations, as well as smaller chromosomal microdeletions and duplications
  62. What is the method of Metaphase FISH?
    For metaphase FISH, cells are arrested in mitosis as for chromosomal banding. They are then fixed using a mixture of acetic acid and methanol and then “dropped” on a glass microscope slide where they are affixed. DNA probes of a few hundred kilobases (kb) in length are used that match regions the chromosomes containing the DNA sequence in question. These probes are directly hybridized with the chromosomes on the slide (hence, the term "in situ" hybridization); immediate detection of the fluorescence signal is possible via fluorescence microscopy
  63. Metaphase FISH
    • Fluorescence in situ hybridization (FISH) of a patient with a deletion of chromosome 22q, the DiGeorge/VCF syndrome. As shown, the VCF probe only hybridizes with one chromosome 22 (red line and arrows). By comparison, the fluorescent green probe is a control probe that hybridizes to both chromosomes.
  64. Advantages of this FISH are:
    • The resolution of FISH is much better than traditional chromosome banding (FISH can resolve 2 megabases (Mb) in length, compared to 6 Mb for chromosomal banding).
    • FISH can be applied to both dividing (metaphase) and non-dividing (interphase) cells.
    • The protocol is technically fairly straightforward.
    • Hybridization with multiple probes enables detection of translocation products. An example is the BCR-ABL1 fusion of t(9;22) in chronic myeloid leukemia.
    • FISH can identify a range of structural abnormalities including deletions, duplications, aneuploidy and the presence of derivative (structurally rearranged) chromosomes.
    • FISH may be used to monitor recurrent or residual disease in bone marrow transplant patients
  65. How can FISH be used for translocation detection?
    Hybridization with multiple probes
  66. What are disadvantages of FISH?
    • Small mutations, including small deletions and insertions as well as point mutations, cannot be identified.
    • Uniparental disomy (inheritance of both copies of a chromosome from the same parent) will be missed because the probe merely detects the presence or absence of a locus or specific portion of a chromosome and not its source.
    • Chromosomal inversions will be missed since a probe can only detect the presence of a specific sequence but not its precise location within the chromosome.
    • Probes are not yet commercially available for all chromosomal regions.
    • The clinician has to choose the correct FISH probe in order to make an accurate diagnosis
  67. What is the mechanism for uniparental disomy?
    • Chromosomal nondisjunction during meiosis leads, after fertilization, to trisomy. Subsequent loss of one chromosome (rescue) could lead to the formation of cells with a chromosome from each parent or cells in which both chromosomes were from the disomic gamete.
  68. When can we use interphase FISH?
    • Interphase FISH is used when dividing cells are not available (eg, in fully differentiated cells or in tissues that have been fixed and embedded in paraffin).
    • It also improves the resolution of FISH probes
  69. How is interphase FISH done?
    • For interphase FISH, cells are tested with FISH probes without cell cycle synchronization. When using intact nuclei (eg, not from paraffin embedded tissue), the cells are harvested using a hypotonic solution, fixed, and again “dropped” on the slides. For FISH on paraffin embedded tissue, thin slices of pathological specimens are cut and affixed to the slides
    • Because chromosomes are only minimally condensed in interphase, this modification of FISH analysis provides the opportunity to hybridize probes at a high resolution (well under 1 Mb, compared to 2 Mb for metaphase FISH). In the interphase nucleus, chromosomal structures cannot be discerned and only the hybridized probe will light up 
  70. What are the advantages of interphase FISH?
    • Higher resolution than metaphase FISH;
    • the ability to perform the test immediately without culturing the cells, which makes it faster;
    • and the applicability to paraffin embedded sections.
    • Interphase FISH is of special value in prenatal diagnosis of various aneuploidy states, such as trisomy 18 or trisomy 21, in which the ability to obtain results rapidly aids in decision making
  71. The major disadvantage of interphase compared to metaphase FISH is that ..................................................................
    • in interphase FISH the chromosomes themselves cannot be visualized.
    • Thus, information cannot be provided regarding overall chromosome number and composition
  72. What are the features of spectral karyotyping?
    • Spectral karyotyping (SKY, also called multicolor FISH) is a FISH technique that accurately identifies the chromosomal origin of all elements in a karyogram (complete chromosome set) using multiple wavelengths of light to generate signals of many colors.
    • A combination of five fluorochromes can be used as probes to "paint" all 22 autosomes, as well as the X and the Y chromosomes, in different colors.
    • The karyogram is analyzed by a computerized spectral imager and the chromosomes are classified based on their particular emission spectra
  73. What are the advantages of spectral karyotyping?
    • complete karyotyping using automated analysis
    • origin of marker chromosomes, small insertions, and complex rearrangements can be inferred through the presence of color-coded chromosomal segments 
  74. What are the disadvantages of spectral karyotyping?
    • Structural rearrangements within a single chromosome will not be detected.
    • The resolution of SKY is relatively low (up to 15 Mb).
    • SKY may be used when a specific abnormality is suspected, but is not applicable as a screening method
    • Expensive
  75. What is Array comparative genomic hybridization?
    • 1) Comparative genomic hybridization (CGH) allows detection of amplifications and deletions of smaller regions of DNA along the lengths of all of the chromosomes. The technique works by comparing the genomic content (or DNA) of a patient (or target) with a normal control individual (or individuals)
    • 2) The resolution of “classical” or metaphase CGH is relatively low (approximately 15 Mb of DNA)
    • 3) Array CGH is a modification of CGH in which the comparator DNA, RNA, or tissue is arrayed on a glass slide or glass beads. There are three basic types of array: bacterial artificial chromosomes (BAC) arrays, oligonucleotide arrays (typically 60 base pairs in length), or single nucleotide polymorphism (SNP) arrays (typically a few nucleotides). Most SNP-based arrays now also include single locus probes as well as SNPs
  76. What are the two type of arrays?
    • Targeted arrays “target” known microdeletion/microduplication syndromes, as well as other known loci of inherited Mendelian disorders (eg, tuberous sclerosis). The first targeted arrays were arrays of around 500 to 600 BACs.
    • Whole genomic arrays cover the entire genome at varying levels of resolution. The first whole genomic arrays were BAC arrays with around 2600 BACs spaced at about 1 Mb throughout the genome (ie, with about 1 Mb resolution). Oligonucleotide and SNP arrays have supplanted the use of BACs. Most laboratories use either oligonucleotide or SNP arrays with an average resolution of about 35 kb throughout the genome
  77. Which arrays can detect CNV?
    Both SNP and oligonucleotide arrays
  78. What arrays can be used to determine whether there is absence or loss of heterozygosity (AOH or LOH) for different regions, or even entire chromosomes in the presence of normal copy number?
    Only SNP arrays
  79. Which type of UPD can be detected by SNP arrays?
    The absence of biparental inheritance can be seen in uniparental disomy (UPD) or in some cancer tissue samples. UPD can be the result of heterodisomy, in which two different homologous chromosomes from the same parent (either maternal or paternal) are present, instead of the normal biparental contribution (one chromosome from each parent). Alternatively, UPD can occur when there are two identical copies of a single parental chromosome (isodisomy). SNP arrays can only detect UPDs secondary to isodisomies
  80. What are the advantages of CGH arrays?
    • dividing cells and tissue culture are not necessary
    • Resolution of the array is dependent upon the type of array used and the average spacing of the “probes” on the array
  81. What are the disadvantages of CGH array?
    • Balanced structural rearrangements (ie balanced translocations, inversions, insertions) will not be detected, because there is no change in copy number
    • Levels of mosaicism (ie copy number changes in some but not all cells) of 20 percent or less will not be detected
  82. What are the methods for genotyping new mutations?
    • Heteroduplex analysis
    • Single strand conformation analysis
    • Automated sequencing- (Whole genome sequencing)
    • Southern and Northern blotting
  83. What is Heteroduplex analysis?
    • Heteroduplex analysis is used to detect point mutations on one strand of a DNA helix.
    • The technique uses denaturation and reannealing of the double stranded target DNA. If complementary single strands re-anneal, they form a perfectly aligned homoduplex. On the other hand, single strands that are not completely complementary because of the presence of a point-mutation in one strand form a heteroduplex.
    • The failure to anneal at all base positions results in the formation of a "bubble" in the newly formed double strand. As compared to normal DNA, DNA with a bubble migrates more slowly during electrophoresis
    • Does Not identify the location of the mutation within the analyzed fragment
  84. Heteroduplex analysis
    This representation of heteroduplex analysis shows how this technique can be used to distinguish between an individual with normal DNA and heterozygous for a gene mutation. An individual's DNA is amplified by polymerase chain reaction (PCR), and the PCR products from both alleles are mixed and denatured (top section). The PCR products are then reannealed, allowing all combinations of DNA pairing. Homoduplexes are generated by the pairing of normal with normal and mutant with mutant DNA; heteroduplexes are generated from the pairing of of normal with mutant DNA. (middle section). The mutation is denoted by an "X".Lane 2: DNA from an individual homozygous for the normal gene forms only homoduplexes, which run faster through the gel.Lane 3: DNA from an individual heterozygous for the gene mutation forms heteroduplexes, which contain bubbles. This makes the DNA run more slowly through the gel.
  85. What is Single strand conformation analysis?
    • Single strand conformation analysis (SSCA) is based on the observation that single-stranded DNA fragments assume unique conformations that depend on their sequence composition when run in a non-denaturing polyacrylamide gel.
    • Migration through the gel thus depends on chain length as well as strand conformation.
    • Before interpretations can be made, the specific pattern of migration in wild-type control samples must be known. This can be achieved by the addition of normal controls in the run.
    • does not identify the location of the mutation within the analyzed fragment
  86. What is automated sequencing?
    • The most direct approach to mutation detection is automated sequencing.
    • A DNA sequence of interest is amplified in the presence of a primer and dideoxynucleotide chain terminators.
    • Either the primer or the chain terminators are fluorescently labeled.
    • As the sequence undergoes electrophoresis in an automated sequencer, a laser beam excites the fluorescent nucleotides, producing a signal that can be compiled into a DNA sequence
  87. What are the advantages of automated sequencing?
    • Advantages of this method include the precise identification of all DNA variations in the target sequence.
    • Thus, the method can be applied to the detection of known mutations as well as to the identification of unknown mutations.
    • For a large number of inherited conditions, new mutations continue to be discovered and add to the understanding of mutation spectra and genotype-phenotype correlations
  88. Whole genome sequencing allows ...................
    identification of mutations in the entire genome, without the investigator having to choose a gene or chromosome region of interest
  89. What is exome sequencing?
    • “exome sequencing,” in which only the ~3 percent of DNA that is transcribed to messenger RNA is sequenced, can be achieved at a lower price than whole genome sequencing.
    • Although this approach will characterize many of the potentially-deleterious mutations that alter protein structure or abundance, exome sequencing is unable to characterize the noncoding, regulatory variants that are thought to contribute to complex traits and is less good at detecting copy number variants
  90. What are the uses of Southern Blotting?
    Southern blotting can be used to detect small mutations as well as large deletions, duplications and gene rearrangements that alter restriction enzyme cleavage sites or the sizes of the resulting pieces of DNA.
  91. How is Southern blotting done?
    • In this technique, genomic DNA is digested with one or more restriction endonucleases and transferred to a membrane.
    • This membrane is hybridized with a single-stranded radioactively labeled probe, under conditions that facilitate double-strand formation between the probe and those fragments on the gel containing the complementary sequence.
    • When an autoradiographic film is exposed to the membrane and developed, the hybridized sequences become visible as bands.
    • The location of each of these bands corresponds to the size of the fragment to which the probe is bound
  92. What is Northern Blotting?
    A similar technique can be applied when RNA is the primary material. With this method, dubbed "Northern blotting", gene transcript size, abundance and expression patterns in different tissues can be analyzed
  93. What are the advantages to blotting?
    • Advantages of these methods are the potential to detect a wide range of mutations and large structural rearrangements.
    • Southern blotting can also be used to detect changes in gene methylation status (which affects sensitivity to restriction nucleotide digestion).
  94. What are the disadvantages for blotting?
    Disadvantages include the requirement for larger amounts of DNA than the other methods mentioned herein, and the labor and time requirements. Generally, this assay can take a week to perform. The use of radioactive materials is both expensive and hazardous. However, nonradioactive methods, such as chemiluminescence, can be substituted