Next-generation sequencing (NGS) is a high-throughput approach to DNA sequencing using massively parallel processing. Parallel processing allows simultaneous sequencing of many DNA fragments which are then assembled using bioinformatics tools to recreate the longer sequences that can be used for various applications, such as whole-genome sequencing, genotyping, investigating gene copy-number variations, transcriptomic and differential expression studies.
The first step in the NGS workflow is collecting the nucleic acid material from the samples that need to be analyzed. Nucleic acid (DNA or RNA) extraction is typically done using pre-made kits. These kits contain buffers and columns that can be used to lyse the cells and isolate DNA or RNA, depending on the sequencing application. Here we will focus on DNA sequencing. For information on RNA sequencing, please refer to our RNA-Seq guide.
It is a good practice to assess the quantity and quality of DNA prior at this point. The collected DNA is then subjected to fragmentation to generate smaller pieces that will be sequenced in parallel. Desired fragment size depends on the sequencing technique used. DNA fragmentation can be done by mechanical methods such as sonication or enzymatically using transposons, restriction enzymes or nicking enzymes. Enzymatic fragmentation can result in higher library yields compared to mechanical shearing, as the latter often leads to sample loss and DNA damage.
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Now that the DNA is fragmented into suitable size pieces, a DNA sequencing library is generated by a Polymerase Chain Reaction (PCR). First, short pieces of synthetic DNA, called adapters, are ligated to the library fragments using DNA ligase. These adapters enable the library fragments to bind the sequencer flow cell and contain the sequencing primer binding sites and indexes. The indexes allow pooling together DNA from different samples and keeping track of which sample the fragments came from during and after sequencing.
The library is now ready for clonal amplification, a process in which each DNA fragment is amplified to create a cluster of identical molecules that enhances the sequencing signal. Two main methods are used for this purpose: emulsion PCR and bridge PCR.
In emulsion PCR, emulsion oil, beads, PCR mix, and the library DNA are mixed, which leads to the formation of microscopic water droplets. Ideally, each droplet has one bead and one DNA molecule. The DNA molecule is denatured to form two separate strands, one of which anneals to the bead. The annealed DNA is amplified by polymerase starting from the bead towards the primer (adapter) site. The original reverse strand is released from the bead only to re-anneal to the bead at a different site to give two separate strands. These are both amplified to give two DNA strands attached to the bead. The process is then repeated at least 30-60 cycles to create clusters of DNA.
Emulsion PCR is inefficient relative to other methods, like bridge PCR, as only a small fraction of droplets, those with only a single bead, are usable.
Bridge PCR uses a flow cell with covalently attached forward and reverse primers. These primers are complementary to the adapters attached to the library fragments, which allows them to attach to the surface of the cell. The two strands are denatured and one of them is washed off. The free end of the single-stranded DNA attaches to the second primer on the flow cell surface , creating bridged structures. Polymerase amplifies the strands, generating a double-stranded bridge, so when denaturation occurs, two single stranded DNA fragments are attached to the surface in close proximity. Repetition of this process leads to formation of clusters of identical strands.
Prior to clonal amplification step, you will need to determine how much of the whole DNA sequence will be sequenced. Non-targeted sequencing amplifies and sequences the entire DNA sample provided, and is useful in whole-genome sequencing. Targeted sequencing utilizes oligos, or primers, to sequence specific areas of interest. This allows for the targeted sequencing of desired regions and may generate a more manageable amount of data to analyze. The most popular methods of targeted sequencing are hybridization capture and amplicon sequencing. Both methods are conducted after DNA fragmentation to enrich fragments containing the sequences of interest.
Not all library preparation involves Polymerase Chain Reaction (PCR) for amplification, like long-read sequencing. There are prefabricated kits with adapters that are synchronized to oligo sequences for the flow cell, such as Illumina sequencing.
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Illumina sequencing platform uses sequencing by synthesis technology. At the end of clonal amplification, the reverse strands are washed off the flow cell, leaving only forward strands. A primer attaches to the forward strands’ adapter and a polymerase adds a fluorescently tagged dNTP** to the DNA strand. Only one base can be added per round due to the fluorophore containing a blocking group. Each of the four bases has a unique emission, and after each round the machine records the color signal emitting from the base added. Once the color is recorded the fluorophore is washed away, together with the blocking group, and another dNTP is added to growing chain, repeating the process.
Sequencing occurs simultaneously for millions of clusters creating a large amount of sequencing data. The data is analyzed by aligning overlapping reads to create continuous sequence stretches, called contigs. If a reference sequence is available, the contigs are compared to it for variant identification. However, contigs are relatively short and may be difficult to align, especially if there are many short tandem repeats or other repetitive sequences.
While Illumina sequencing is a common method of high-throughput sequencing, other sequencing methods exist and have their own pros and cons. Ion Torrent sequencing detects the release of hydrogen when a complementary dNTP pairs forms with the target DNA strand. This process is repeated with each dNTP to sequence each strand. This process is fast and low cost but has difficulty detecting long repeats of the same base (e.g. AAAAAAA) and produces shorter read lengths relative to other methods. Nanopore sequencing uses electrophoresis to transport an unknown sample through a nanopore. Sequencing occurs as the sample passes through because each base pair alters the density of the electric current flowing through the nanopore, which can be detected and analyzed by software. Nanopore sequencing can sequence small amounts of DNA with few reagents but tends to be highly error-prone.
**The function of Deoxynucleoside triphosphate (dNTP)s in PCR is to expand the growing DNA strand with the help of Taq DNA polymerase. It binds with the complementary DNA strand by hydrogen bonds. The PCR is an in vitro technique of DNA synthesis
1. Developing a primary culture
Primary culture refers to the cells that are isolated directly from the tissue of interest and proliferated until they reach confluence, or occupy all the available substrate. One method of acquiring cells for primary culture entails sampling from the tissue directly. Cells taken in this manner must be disaggregated using enzymatic or mechanical means before they are placed on the substrate.
After the primary culture reaches confluence, the cells have to be subcultured by transferring them to fresh growth medium. At this stage the cell culture is no longer considered primary and becomes secondary culture, or cell line. Cell lines derived from primary cultures are finite, which means they have a limited number of cell divisions that is a genetically determined. The loss of ability to proliferate is called senescence.
Another way of acquiring cell culture is by using an established cell strain. These are continuous, or immortalized, cell lines that have been mutated and lost the ability to undergo senescence. These cells can continuously divide and are optimal cell lines for prolonged studies. The most common immortalized cell lines used for primary culture include HeLa cells and HEK 293 cells, among others
2. Developing Cell Lines
Regardless of the cell line type, it is necessary to propagate the cell line by continuously passaging the cells into fresh media, a process known as splitting the cells. To split cells, cell media is aspirated via a vacuum and the cells are washed in warmed Phosphate Buffered Saline (PBS) to remove residual media. If the cells are anchorage-dependent and rely on adherence to a surface, the cells can be treated with trypsin, a chemical that causes cells to temporarily lose their adherence. Trypsin, however, is toxic and should not be applied for long.
Trypsin toxicity is neutralized by adding growth media. For mammalian cells, this growth media is typically DMEM supplemented with Fetal Bovine Serum (FBS) and antibiotics. From this solution, the media and cells can be partitioned, or split, into new cell culture plates. A common practice is to always maintain at least one plate for further splitting and use the rest to split into additional plates that will be used for experiments.
While the above example is for anchorage-dependent mammalian cells, a similar principle applies to other types of cells including those in suspension and spore colony culture. In suspension culture, cells from a developed culture may be split by transfer into fresh growth media. In colony culture, spores or colonies may be taken from a developed plate and streaked across fresh media plates
Different cells require different growth conditions. While conditions may vary from cell type to cell type, most cells are grown in incubators to maintain optimum growth conditions. Incubators have the ability to maintain an optimum temperature (37oC for most cells) and precise O2 and CO2 levels. Additionally, cells will require a growth medium that is unique to each cell type. Media contains the nutrients, hormones, and buffers for cells to grow and should be continually changed. The right media and incubation conditions are critical for cell growth.
3. Maintaining Cell Lines
Successful cell culture depends on keeping the cells free from contamination by microorganisms such as bacterial, fungi, and viruses. Nonsterile supplies, media, reagents, airborne particles, unclean incubators, and dirty work surfaces are all sources of biological contamination.
Aseptic technique is the best barrier to prevent contamination by invasive microorganisms and should always be maintained. Aseptic technique includes:
- Only opening cell culture in a sterile environment such as a cell culture hood with working airflow.
- Sterilizing the work area before and after each handling of the cells with proper cleaning reagents. 70% ethanol is the most common choice.
- Using gloves, lab coats, and other PPE as needed to prevent contamination of the samples and maintain personal safety.
- All media and solutions should be opened and/or mixed within a cell culture hood.
- Using only sterile glassware and disposable pipettes. Dispose of pipettes after each use.
- Opening media and other containers only when ready to use.
However, even with proper aseptic technique, contamination can happen. In these cases, the incubator and cell culture hood should be thoroughly cleaned to prevent other samples from being contaminated. If contaminated cells or media are critical to work, they may be treated with antibiotics to kill the contaminants - though care should be taken as antibiotics may cross-react with the cells of interest. In most cases, the contaminated cells and media should be thrown out and prepared anew.
Long-term storage of cells is a useful way to backup any experiments relying on a certain subculture or strain of cells. The most common method is Cryopreservation. Cryopreservation requires a surplus of cells be taken from cell culture and mixed with a protective agent, typically DMSO or glycerol, before being stored below –130°C.