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Over the course of six decades, large amounts of time and resources have been invested in developing and improving the technologies that underpin genetic research. DNA sequencing is one method that has seen a vast improvement over the years. When considering the history of this technique, researchers have gone from being able to sequence only a short oligonucleotide of a single gene to the whole genome sequencing (WGS) that is available now. DNA sequencing can be summarised in two generations from the genesis of this field to the starting time of this study.

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1.10.1 First-generation DNA sequencing

Over two decades elapsed from the discovery of the double helix structure of DNA (Watson and Crick, 1953) to the introduction of several influential DNA sequencing protocols. Initial efforts at sequencing were met with limited success since the methods employed could only determine the nucleotide composition but were not powerful enough to determine the order of the nucleotides (Holley et al., 1961). However, the mid-1970s represent the real start of ‘first-generation’ DNA sequencing by widely adopted the plus and minus and the chemical cleavage techniques (Sanger and Coulson, 1975; Maxam and Gilbert, 1977). This method used radio-labelled DNA treated with chemicals designed to cleave fragments of the chain at specific bases, followed by migrating the labelled fragments through a polyacrylamide gel to determine the length and position of the nucleotides.

Sanger's ‘chain-termination’ or dideoxy technique (Sanger et al., 1977) represented the most significant development amongst the first generation of DNA sequencing methods. This technique used polymerase-based copying of single-stranded DNA, but included a small proportion of radio-labelled chemical analogues of the nucleotides, chain-terminating dideoxynucleotides (ddNTPs), in each of four parallel reactions. The products were then run in adjacent lanes on a polyacrylamide gel to produce radioactive bands in the lanes, the positions of which corresponded to the sequence of nucleotides. Over time further improvements were made to this technique. Fluorescent dyes (Smith et al., 1986) were used instead of radioactive labelling to tag the different chain terminating analogues, allowing all four nucleotides to be resolved in a single lane. Capillary electrophoresis instead of a polyacrylamide gels, together with the use of laser induced fluorescence detection, allowed longer reads and avoided the need to cast new gels for each sequence. (Ruiz-Martinez et al., 1993; Hebenbrock et al., 1995). The polymerase chain reaction (PCR) replaced DNA cloning from libraries as the main method to generate the sequencing templates (Saiki et al., 1985; Saiki et al., 1988) and automated Sanger sequencing by capillary electrophoresis was established (Hunkapiller et al., 1991). This allowed 500-1000bp of DNA sample to be sequenced in 6–8 hours. This method is the gold-standard DNA sequencing technique that is still used in laboratories today to sequence short pieces of DNA.

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1.10.2 Second-generation DNA sequencing

The emergence of a new technique known as pyrosequencing (Roche GS FLX) paved the way for ‘next-generation sequencing’ (NGS) technologies for high-throughput sequencing. In this method, an enzymatic reaction occurs in which ATP sulfurylase converts pyrophosphate into ATP, which subsequently serves as the substrate for luciferase, meaning that the light produced is proportional to the amount of pyrophosphate (Nyrén and Lundin, 1985). The amount of incorporation is monitored by luminometric detection of the quantity of pyrophosphate released as each nucleotide is washed through the system in turn over the template DNA affixed to a solid phase. This signal is then used to infer DNA sequences (Hyman, 1988). The weakness with this technique though, is that errors can be caused by misjudging the length of homopolymer runs in this process, which may result in false single-base insertions or deletions (indels) in the DNA sequencereadout (Ronaghi et al., 1998).

The sequencing machines developed by different companies during the first decade of the new millennium dramatically increased the amount of DNA that could be sequenced, ranging from five hundred million bases of raw sequence (Roche) to billions of bases in a single run (Illumina, ABI SOLiD technology). These machines relied on performing massive numbers of parallel sequencing reactions on a micrometer scale on clonal beads (Roche and ABI SOLiD) or clonal bridges (Illumina). (Shendure and Ji, 2008). Over the last decade, these three platforms are commercially leading second generation NGS platforms (Pareek et al., 2011).

Here at Leeds the NGS facility (a partnership between the University of Leeds and the Leeds Teaching Hospitals NHS Trust) uses the Illumina NGS platform. For the Illumina Genome Analyser, sample preparation involves fragmentation of the DNA sample, enzymatically repairing the staggered ends, adding adenines (A) to the 3-ends of the DNA fragments and ligating adapters, followed by library amplification (Myllykangas et al., 2012). Solid-phase amplification is used to produce randomly distributed, clonally amplified clusters from fragments or mate-pair templates on a glass slide. The sequencing library is immobilised on the surface of a flow cell onto which a “lawn” of high-density forward and reverse primers has been covalently attached to the slide to create an ultra-dense primer field. The primer functionalised flow cell surface

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serves as a support for amplification of the immobilised sequencing library by a process also known as “Bridge-PCR” (Figure 1.11A). The resulting bridged double-strand DNA is freed using a denaturing reagent. Repeated reagent flush cycles generate groups of thousands of DNA molecules, also known as “clusters,” on each flow cell lane. DNA clusters are then unbound from the complementary DNA strand (linearization), followed by blocking the free 3’-ends of the clusters and hybridising a sequencing primer.

Figure 1.11. Illumina solid-phase amplification and four-colour cyclic reversible termination sequencing method. Illumina solid-phase amplification (A) is achieved through two

basic steps, these being initial priming and extending of the single stranded single-molecule template, followed by bridge amplification of the immobilised template with immediately adjacent primers to form clusters. The four-colour cyclic reversible termination (B) uses Illumina’s 3′-O-azidomethyl reversible terminator chemistry on solid-phase-amplified template clusters. Following incorporation, a cleavage step removes the fluorescent dyes and regenerates the 3′-OH group using the reducing agent Tris (2-carboxyethyl) phosphine. (C) The four-colour images highlight the sequencing data from two clonally amplified templates. (Adapted from Metzker (2010) with the permission of Elsevier Copyright Clearance Centre, License number: 3940451410070).

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Cyclic reversible termination is the method employed by Illumina for sequencing- by-synthesis. Firstly, a DNA polymerase bound to the primed template incorporates just one fluorescently modified nucleotide, which represents the complement of the template base. Following incorporation, the remaining unincorporated nucleotides are washed away (Figure 1.11B). Secondly the four colours of the four nucleotides are detected by total internal reflection fluorescence imaging using two lasers (Figure 1.11C). The synchronous extension of the sequencing strand by one nucleotide per cycle ensures that homopolymer stretches can be accurately sequenced. However, failure to incorporate a nucleotide during a sequencing cycle results in an off-phasing effect, and as the sequence extends, gradually more and more molecules lag behind in the extension so that the generalised signal derived from each cluster deteriorates over many cycles. Therefore, Illumina sequencing accuracy declines as the read length increases, which limits this technology to short sequence reads (Myllykangas et al., 2012).