DNA extraction kits

We are currently using two different types of DNA extraction kits, depending essentially on the quantity of DNA required. The first – the Qiagen system – is a robust method for obtaining ≈ 5 μg of DNA from 200 μl of whole blood ( www.qiagen.com ). This method utilises a high-affinity DNA binding matrix in a spin column, which can be used in a microcentrifuge or in an automated platform, such as the Qiacube. The DNA obtained is generally of high quality and sufficient in amount for most routine analyses.

When a larger amount of DNA is required – perhaps for storage and more extensive analysis of critical samples – we use from 3–10 ml of blood with the Gentra Puregene blood extraction kit (Qiagen), which can yield upward of 100 μg of DNA. This method depends on a salting-out of proteins after sequential red cell and then white cell lysis, followed by an isopropanol precipitation of the DNA. Other nucleic acid high throughput systems also exist such as MagNA Pure 96 System (Roche, http://lifescience.roche.com ) and the Maxwell® rapid sample concentrator (RSC) Instrument ( www.promega.com ).

Protocols will be provided in the latter part of this chapter. For preparation of reagents and a protocol for the manual extraction of DNA from blood using organic solvents, the reader is referred to previous editions of this book.

Principle

A DNA polymerase will synthesise the complementary strand of a DNA template in vitro. A stretch of double- stranded DNA is required for synthesis to be initiated. This double-stranded sequence can be generated by annealing an oligonucleotide (‘oligo’), which is a short, single- stranded DNA molecule usually between 18 and 25 bases in length, to a single-stranded DNA template. These oligos, which are synthesised in vitro, will prime the DNA synthesis and are therefore referred to as ‘primers’.

In PCR, two oligo-primers are used. Each primer will generate a copy of the complementary strand. For instance, the sense primer will anneal to the antisense sequence and prime it in a 5′ to 3′ direction and in doing so generate a new sense strand. The antisense primer will anneal to the sense strand and prime it generating a new antisense strand. The other components of the reaction are the DNA template from which the DNA fragment will be amplified, the four deoxynucleotide triphosphates (dATP, dTTP, dCTP and dGTP) required as the building blocks of the newly synthesised DNA, a salt buffer containing MgCl ₂ and the thermostable DNA polymerase (Taq polymerase).

The first step of the reaction is to denature the DNA, generating single-stranded templates, by heating the reaction mixture to 95 °C. The reaction is then cooled, usually to a temperature between 50 °C and 68 °C, which permits the annealing of the oligos to the DNA template but only at their specific complementary sequences. The temperature is then raised to 72 °C, at which temperature the Taq polymerase efficiently synthesises DNA, extending from the oligo in a 5′ to 3′ direction. Cyclical repetition of the denaturing, annealing and extension steps, by simply changing the temperature of the reaction in an automated heating block, results in exponential amplification of the DNA that lies between the two primers ( Fig. 8-1 ). So-called fast-cycling Taq kits are now available, often with highly modified cycling conditions that can bring the time of a PCR reaction down to below an hour.

The polymerase chain reaction.

Cyclical repetition of three temperatures for denaturing, annealing and extending DNA, respectively, gives rise to an exponential amplification of a DNA fragment between two primer sequences directing DNA synthesis on opposite strands of the DNA template.

The specificity of the DNA fragment that is amplified is therefore determined by the sequences of the primers used. A sequence of 18–25 base pairs (bp) is theoretically unique in the human genome, and so primers of this length and longer will anneal at only one specific place on a template of genomic DNA, or at least one combination within an amplifiable distance (typically no more than 2–3 kilobases for standard cycling conditions). One general requirement of PCR is therefore some knowledge of the DNA sequence of the gene that is to be amplified. The relative positioning of the two primers is another important consideration. They must prime DNA synthesis in opposite directions but pointing toward one another and each must anneal to one of the complementary strands. There is also an upper limit to the distance apart that the oligos can be placed; fragments of several kilobase (kb) pairs in length can be amplified, but the process is most efficient for fragments of several hundred base pairs. Long-range PCR kits are available, which in combination with optimised cycling conditions can amplify fragments of 10–20 kb.

Methodology

Modifications and developments

The procedure described above is a guide for setting up and checking a standard PCR amplification. As the test dictates, modifications can be used, such as the following:

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    Radiolabelling. A PCR can be labelled with ³² P by adding 0.1 μl of [α- ³² P]dCTP per tube to the reaction mixture (rarely used).

    
    
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    Multiplex. More than one fragment can be amplified in the same tube simply by adding in further primer pairs. It is important that the different pairs all work equally well under the same conditions.

    
    
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    Nested PCR. This involves successive rounds of amplification using two pairs of primers; the second pair, located within the sequence amplified by the first, allows products to be generated from as little as a single cell.

    
    
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    Long-range amplification. Fragments upward of 10 kb can now be generated by PCR using modified polymerases.

    
    
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    Automation. High throughput PCR amplification is being achieved through the use of robots and 96-well plate technology.

    
    
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    Automated fragment analysis. The method of gel electrophoresis is modified for the detection of fluorescently labelled PCR products on DNA fragment analysers (e.g. the ABI 3700 DNA analyser, www.thermofisher.com ).

  
  

Interpretation

If the amplification has been successful, a discrete fragment of the expected size is seen in a SYBR Safe-stained agarose gel in all samples, except in the NTC lane. If a product is seen in the NTC, then one of the solutions has been contaminated and the results cannot be relied on. All the working solutions must be discarded and the micropipettes must be cleaned. Cleaning micropipettes and work surfaces prior to the start of each run is highly recommended. To avoid contamination, making up a stock master mix of all reagents is recommended before DNA samples are added to each tube. If possible the use of separate pre- and post-PCR laboratories is highly recommended. This reduces the risk of contamination significantly. At a minimum the use of distinct areas of the laboratory for each activity is recommended.

Problems

The absence of a fragment in all tracks, including the positive control, indicates that the PCR has failed. This can occur for a number of reasons, including poor quality template or omission of one of the essential reagents. The reaction may also fail if the magnesium concentration is too low (standard concentration 1.5 mM) or if the annealing temperature is too high. However, DNA quality and amount in the reaction is often one of the major reasons for failure. If one particular DNA sample repeatedly fails to amplify, then the sample should be re-extracted using Proteinase K ( www.thermofisher.com ) and phenol/chloroform and re-precipitated, adding in one-tenth of the total volume of 3 M sodium acetate (pH 4.8) followed by the addition of 2.5 volumes of − 20 °C cold ethanol.

We have also found that using the Gentra blood extraction kit (Qiagen) or passing the DNA through a Qiagen column substantially improves DNA quality and PCR efficiency.

Another problem is the presence of nonspecific fragments or just a smear of amplified product. This can occur if the magnesium concentration is too high or if the annealing temperature is too low.

Principle

Restriction enzymes (REs) cleave DNA at short specific sequences. Because many REs are available, it is not uncommon for a single point mutation to coincidentally create or destroy an RE recognition sequence. If this is the case, digestion of the appropriate PCR product prior to agarose gel electrophoresis enables the mutation to be identified. A difference in the size of the restriction fragments seen in normal and mutant samples can be predicted from a restriction map of the amplified fragment and the site of the mutation that changes a restriction site. The observed fragments should be consistent with either the mutant or the normal pattern. An example of this technique is shown in Figure  8-2 .

Detection of the sickle cell mutation using restriction enzyme analysis.

A, A sketch of the β globin gene shows the position of the primers used to amplify a 768 bp fragment in a polymerase chain reaction (PCR). The sequence of β I–5′ is

5′ TAAGCCAGTGCCAGAAAGAGCC 3′ and that of β II-3′ is

5′ CATTCGTCTGTTTCCCATTCTA 3′. Maps of the Bsu36 I restriction sites and the fragment sizes from βA and βS genes are shown below. B, An ethidium bromide–stained minigel illustrates the fragment sizes generated by Bsu36 I digestion of the PCR product from normal (A/A), sickle cell trait (A/S) and sickle cell anaemia (S/S) individuals, along with the undigested amplified fragment (U) and the molecular size marker (M).

Methodology

Principle

Under appropriate conditions, short oligonucleotide probes will hybridise to their exact complementary sequence but not to a sequence in which there is even a single base mismatch. A pair of oligos is therefore used to test for the presence of a point mutation: a variant oligo complementary to the variant sequence and a reference oligo complementary to the reference sequence, with the sequence difference placed near the centre of each oligo.

The stability of the duplex formed between the oligo and the target DNA being tested (the PCR product) depends on the temperature, the base composition and length of the oligo, and the ionic strength of the washing solution. For allele-specific oligonucleotide hybridisation (ASOH) studies, an empirical formula has been derived for the dissociation temperature ( T _d ), the temperature at which half of the duplexes are dissociated. This value is used as a guide; the exact temperature at which only perfect base pairing is maintained is usually determined by trial and error.

This methodology has been widely applied for the detection of single nucleotide variations (SNVs) using fluorescently labelled hydrolysis probes (such as TaqMan probes) that distinguish the two alleles. Two short allele-specific probes are used, one of which will hybridise only to the wild-type allele and one of which will hybridise only to the mutant allele. The two probes are labelled with different fluorophores, which are quenched (preferably using a non-fluorescent quencher) while the probes remain intact, but are released if and when the probe hybridises to its perfectly complementary sequence during the reaction, as it will then be broken up by the exonuclease activity of the Taq polymerase.

Interpretation

The oligos will hybridise to their perfectly complementary DNA sequence, such that the variant oligo gives a signal only when the mutant allele is present, and similarly for the wild-type allele. When this is the case, the interpretation of the result is straightforward; a positive signal from a particular oligo indicates the presence of that allele in the test sample. Heterozygotes and homozygotes are distinguished by using the variant and reference oligos in tandem. With the two fluorescent colours of the hydrolysis probes, the heterozygote is identified as the sum of the two fluorochromes.

Other non-radioactive probes, with detection systems involving horse-radish peroxidase, have also been quite widely used in this procedure. The technique has also been modified such that the allele-specific oligonucleotides are immobilised onto nylon membranes and the patient-specific PCR product is used as the probe – the reverse dot blot procedure. This allows several different mutations to be analysed simultaneously and has proved particularly useful in the diagnosis of β thalassaemia mutations.

Sickle cell disease

The presence of haemoglobin S and thus the presence of the sickle cell gene can be determined by haemoglobin cellulose acetate electrophoresis, high performance liquid chromatography (HPLC) or a ‘sickling test’. However, there are occasions when it is beneficial to make a diagnosis by DNA analysis (e.g. in prenatal diagnosis, which can be performed at 10 weeks of pregnancy, with the aim of identifying sickle cell anaemia and haemoglobin S/β thalassaemia, or to confirm the diagnosis of sickle cell anaemia in a neonate). For the type of specimens collected for prenatal diagnosis, refer to page 310.

The sickle cell mutation, c.20A > T; p.Glu7Val in codon 7 of the β globin gene, HBB, results in the loss of a Bsu36 I (or Mst II, Sau I, OxaN I or Dde I) restriction enzyme site that is present in the normal HBB gene. It is therefore possible to detect the mutation directly by restriction enzyme analysis of a DNA fragment generated by PCR. A pair of primers are used to amplify exons 1 and 2 of the β globin gene and the products of the PCR are digested with Bsu36 I. The loss of a Bsu36 I site in the sickle cell gene gives rise to an abnormally large restriction fragment that is not seen in normal individuals ( Fig. 8-2 ). This mutation can also be detected by ARMS PCR using two reactions per sample, one specific for the mutant allele and the other for the wild-type allele. The 3′-end nucleotide of the mutation-specific primer should be specifically complementary to the mutation, while the primer specific for the wild-type sequence should contain a nucleotide complementary to the wild-type sequence at this end. A common reverse primer allows the production of an amplification band similar in size in the two reactions, hence the use of two reaction tubes per sample. Primers specifically used for this analysis are listed below.

Homozygosity for the mutation will produce amplification in the mutation PCR reaction, while the absence of the mutation will result in amplification in the wild-type PCR only. Heterozygous samples will have amplification in both reactions. An extra pair of primers amplifying another part of the gene should be used to produce control bands. The primers used for this method are Haemoglobin S Forward, Wild-Type (WT) Forward, Common Reverse, Control Forward, and Control Reverse and are listed here: Haemoglobin S Forward Mutant: 5′-CCC ACA GGG CAG TAA CGG CAG ACT TCT GCA 3′; Haemoglobin S Forward WT: 5′-CCC ACA GGG CAG TAA CGG CAG ACT TCT GCT 3′; Common Reverse: 5′-ACC TCA CCC TGT GGA GCC AC 3′; Control Forward: 5′-GAG TCA AGG CTG AGA GAT GCA GGA 3′; Control Reverse: 5′-CAA TGT ATC ATG CCT CTT TGC ACC 3′.

β thalassaemia

The ethnic groups with the highest incidence of β thalassaemia are the Mediterranean populations, Indian subcontinent populations, Chinese and Africans. Although more than 100 β thalassaemia mutations are known, each of these groups has its own subset of mutations, so that as few as five different mutations may account for more than 90% of the affected individuals in a population. This makes the direct detection of β thalassaemia mutations a reasonable possibility, and it has become the method of choice where it is most important – in prenatal diagnosis. ^,

The majority of mutations causing β thalassaemia are point mutations affecting the coding sequence or splice sites and nowadays they are detected by direct DNA sequence analysis. Unstable and other unusual haemoglobins may also cause disease, and can similarly be identified by direct DNA sequence analysis. An example of such a case is shown in Figure 8-3 , where a picture of moderate anaemia is seen in the heterozygote due to the highly unstable and electrophoretically silent variant HBB Durham-N.C.

Sequencing of the β globin gene reveals haemoglobin Durham-N.C.

The sequencing electrophoretogram clearly shows the heterozygous mutation (arrowed) where a C peak and a T peak are superimposed and called as N. This mutation is c.344 T > C causing a p.Leu115Pro substitution (traditionally named as codon 114).

(Courtesy A. Tibepu.)

α thalassaemia

In contrast to the β thalassaemias, the most common α thalassaemia mutations are large deletions. Two categories exist: those that remove only one of the two α globin genes on one chromosome (α ⁺ thalassaemia) and those that remove both of the α genes from one chromosome (α ⁰ thalassaemia). Although PCR amplification around the α globin locus has proved to be rather difficult, the common deletions can now be identified by a reasonably robust Gap-PCR. In these reactions dimethyl sulphoxide (DMSO) (see Buffer II, p. 128 ) and betaine are added. Two different multiplex PCR reactions are set up: one for the common α ⁺ thalassaemias (α ^{− 3.7} and α ^{− 4.2} ) and one for the common α ⁰ thalassaemias (– – ^SEA , – – ^MED , – – ^FIL and α ^{− 20.5} ). The fragment generated by these primers across the deletion breakpoint is different in size from the fragment generated from the normal chromosome. The primers that flank the deletion breakpoint are too far apart to generate a fragment from the normal chromosome in the PCR. Only when these are brought closer together as a result of the deletion, can a fragment be produced. Primer sequences used in this analysis are given in Table 8-1 and an example of their application in the detection of α ⁰ thalassaemias is shown in Figure 8-4 .

Detection of α ⁰ thalassaemia by multiplex Gap-PCR.

The sequences of the primers used are shown in Table 8-1 . A normal fragment of 1010 bp is generated by the primers α/SEA(F) and α/(R) in all lanes (although this is very faint in lane 11). In addition, a fragment of 660 bp is generated by the primer pair α/SEA(F) and SEA(R) in lanes 1, 4 and 8 in individuals who are heterozygous for the – – ^SEA deletion; a fragment of 550 bp is generated by the primer pair FIL(F) and FIL(R) in lane 9 in an individual who is heterozygous for the – – ^FIL deletion; a fragment of 875 bp is generated by the primer pair MED(F) and MED(R) in lane 10 in an individual who is heterozygous for the – – ^MED deletion; and a fragment of 1187 bp is generated by the primer pair 20.5(F) and 20.5(R) in lane 10 in an individual who is heterozygous for the –α ^20.5 deletion.

Recently, with the development of new and better PCR reagents, it has been possible to perform a multiplex PCR for the detection of the common α globin gene deletions in one reaction tube. These reagents do not need the addition of DMSO or betaine. More than 30 non-deletional forms of α thalassaemia have been described. Of these, haemoglobin Constant Spring and the α ^HphI mutation are relatively common in Southeast Asian and Mediterranean populations, respectively. These can be detected by ASOH, ARMS, restriction enzyme digestion or direct sequencing of the appropriate PCR product. Unlike the β thalassaemias, α thalassaemias are not easily diagnosed using routine haematological techniques. The diagnosis of α thalassaemia is often made following exclusion of β thalassaemia and iron deficiency. Because the vast majority of cases of α thalassaemia are of the clinically benign type (i.e. α ⁺ thalassaemia), it is debatable whether molecular analysis is justified to reach a diagnosis in these individuals. However, it is important that individuals with α ⁰ thalassaemia are identified and the only definitive diagnostic test is DNA analysis. The α ⁰ thalassaemias are almost entirely restricted to certain ethnic groups, namely, those of Chinese, Southeast Asian and Mediterranean origin; and so it is most efficient to target these groups specifically. The diagnosis of α ⁰ thalassaemia is particularly relevant if prenatal diagnosis is to be offered to a couple who are at risk of having a foetus with hydrops, since there is an increased risk of maternal death at delivery. Guidelines derived from the UK experience as to how and when DNA analysis should be implemented are available. Primers used for this analysis are listed in Table 8-1 and an example is illustrated in Figure 8-4 .

Thrombophilia screening

Considerable advances have been made in our understanding of the genetic risk factors found in patients with venous thromboembolism (VTE). Among these are the diverse mutations causing protein C, protein S and anti-thrombin deficiency. An increased factor VIII level is also a risk factor for VTE, but the genetic determinants of this are unclear. Homozygosity for the common C677T mutation of the methylenetetrahydrofolate reductase gene (MTHFR), which gives rise to a thermolabile variant of this protein, has been reported to be a risk factor for VTE, although other studies have not supported this claim and we no longer provide this test as it appears to have no clinical impact.

A point mutation in the 3′ UTR of the prothrombin gene ( FII ) associated with elevated protein levels has been identified as a genetic risk factor for VTE. The most common of the known genetic risk factors for VTE is a resistance to the anticoagulant effect of activated protein C caused by the Arg506Gln substitution in the factor V gene ( FV ). Around 20% of subjects of north European origin presenting for the first time with thromboembolism are heterozygous for this mutation, designated factor V Leiden (FVL). Because of their prevalence and because the tests have become relatively simple, there is a tendency toward indiscriminate testing for these genetic risk factors in thrombophilia, but without careful and informed counselling this may often be inappropriate. (See also Chapter 19 .)

Interpretation

The results can be evaluated using the allelic discrimination plots for the individual genotyping test. Each data point represents one sample and they can be seen to fall into four clusters according to the genotype of the sample or the absence of the signal (NTC) ( Fig. 8-5 ). If the assay has worked, these clusters should be clearly distinct. An automatic genotype annotation is then based on this clustering. Some points will not be automatically identified by the software: these data points can be annotated manually by assigning them to the appropriate cluster.

Allelic discrimination of Factor V Leiden using the TaqMan assay system.

Each spot on the graph represents one well of the plate in which one DNA template is under test. In each well we either see no signal (negative: bottom left) – NTC; only ‘red’ signals, coming from the wild-type probe (bottom right) – homozygous wild-type; only blue signals coming from the FVL probe (top left) – homozygous FVL; or a mixture of signals to give green spots coming from both the wild-type and FVL probes (top right) – heterozygous for FVL.

Clotting disorders

Diverse mutations underlie haemophilia A and haemophilia B and these are usually identified in specialised laboratories by screening exons for mutation by single-strand conformation polymorphism analysis (SSCP), denaturing HPLC or direct DNA sequence analysis. It may still be relevant to determine carrier status and offer prenatal diagnosis through genetic linkage analysis. Problems with this include the number of sporadic cases, lack of informative markers, unavailable family members and the possibility of recombination.

Of particular diagnostic significance is the fact that from between one third and one half of all patients with severe haemophilia A have a large genomic inversion mutation involving recombination between a region in intron 22 of the factor VIII gene and telomeric homologous sequences. These inversions are readily detected by Southern blot analysis applying the p482.6 probe to Bcl I digests of genomic DNA. A method has also recently been described using long-distance PCR, enabling identification of these deletion mutations in a single tube reaction. (See Chapter 18 for more information on bleeding disorders.)

Cytogenetic analysis

Principles and Terminology

Cytogenetic analysis is usually carried out by specially trained scientists in a separate laboratory that often has no specific relationship to the routine haematology laboratory. For this reason no details of techniques will be given. However, cytogenetic analysis is so crucial to the diagnosis and management of haematological neoplasms that it is necessary for haematologists to understand the principles and be able to understand the reports that are received. In addition, haematologists are often involved in collection of appropriate samples.

Classical cytogenetic analysis is carried out on cells that have entered mitosis and have been arrested in metaphase so that individual chromosomes can be recognised by their size and their banding pattern following staining (e.g. Giemsa staining [G-banding] or staining with a fluorescent dye). Alternating dark and light bands are numbered from the centromere toward the telomere to facilitate description of any abnormalities detected. An example showing the balanced translocation t(9;22)(q34;q11.2) in CML is shown in Figure 8-6 . The standard terminology applied to chromosomes is shown in Table 8-2 .

G-Banded karyotype.

Female karyotype with a balanced reciprocal translocation between chromosomes 9 and 22 (arrowed) giving rise to the Philadelphia (Ph) chromosome.

(Courtesy J. Howard.)

Table 8-2

Terminology and abbreviations used in classical cytogenetic analysis

Term	Abbreviation	Explanation
Centromere	cen	The junction of the short and long arms of a chromosome
Telomere	ter	The termination of the short or long arm of a chromosome, pter or qter
Long arm	q	The longer of the two arms of the chromosome that are joined at the centromere
Short arm	p	The shorter of the two arms of the chromosome that are joined at the centromere
Diploid		Having the full complement of 46 chromosomes, 44 paired autosomes, and two sex chromosomes in a cell or clone
Haploid		Having 23 chromosomes, a single copy of each autosome, and either an X or a Y chromosome in a cell or clone
Tetraploid		Having a total of 92 chromosomes, four of each autosome and four sex chromosomes in a cell or clone
Aneuploid		Having a chromosome number that is neither diploid nor a fraction or a multiple of the diploid number, in a cell or clone
Pseudodiploid		Having 46 chromosomes in a cell or clone but with either structural abnormalities or with loss and gain of different chromosomes so that not all chromosomes are paired
Hyperdiploid		The presence of more than 46 chromosomes in a cell or clone
Hypodiploid		The presence of fewer than 46 chromosomes in a cell or clone
Monosomy	− (a minus sign before the chromosome number, e.g. − 7)	Loss of one of a pair of chromosomes
Trisomy	+ (a plus sign before the chromosome number, e.g. + 13)	Gain of a chromosome so that there are three rather than two copies
Deletion	del or a minus sign after the number and the designation of the arm of a chromosome, e.g. del(20q) or 20q −	Loss of part of the long or the short arm of a chromosome
Translocation	t	Movement of a chromosomal segment or segments between two or more chromosomes; a translocation can be reciprocal or nonreciprocal
Reciprocal translocation		Exchange of segments between two or more chromosomes
Nonreciprocal translocation		Movement of a segment of a chromosome from one chromosome to another but without reciprocity
Balanced translocation		A translocation that occurs without loss of chromosomal material, or at least without loss of sufficient chromosomal material to be detectable by microscopic examination of chromosomes
Unbalanced translocation		A translocation that is associated with gain or loss of part of a chromosome
Inversion	inv	The inversion of a part of a chromosome, either pericentric or paracentric
Pericentric inversion		An inversion that follows breaking of both the long and short arms so that the part of the chromosome that is inverted includes the centromere
Paracentric inversion		An inversion that follows the occurrence of two breaks in either the long or the short arm of a chromosome so that the part of the chromosome that is inverted does not include the centromere
Insertion	ins	The insertion of a segment of one chromosome into another chromosome or into a different position on the same chromosomes. Can be direct or inverted
Isochromosome	i	A chromosome with two long arms or two short arms joined at the centromere, e.g. i(17q)
Derivative	der	A chromosome that is derived from another; a derivative chromosome derived from two or more chromosomes carries the number of the chromosome that contributed the centromere
Duplication	dup	The duplication of part of a chromosome
Clone		A population of cells derived from a single cell; in cytogenetic analysis a clone is considered to be present if two cells share the same structural abnormality or extra chromosome or if three cells have lost the same chromosome
Marker	mar	An abnormal chromosome of uncertain origin that ‘marks’ a clone
Constitutional	c	A chromosomal abnormality that is part of the constitution of an individual rather than being acquired, e.g. + 21c in Down syndrome

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