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What determines the amount of data you would get from a sequencing experiment?

What determines the amount of data you would get from a sequencing experiment?


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When someone performs a sequencing experiment, what determines how much sequencing data you get from the machine (Illumina, PacBio, Nanopore, etc). Or maybe, put differently, how does the machine know that it is done?


On an Illumina machine, the instrument runs as many cycles as you tell it to. If you tell it to run 150 cycles, you will get 150-mers as output.


Sequencing Basics

Automated fluorescent sequencing utilizes a variation of the Sanger chain-termination protocol developed over 20 years ago. In this method, the DNA to be sequenced acts as a template molecule to which a short, complementary oligonucleotide (primer) will anneal to begin enzymatic extension and amplification of a specific region of double-stranded DNA. The newly created fragments will be complementary to the template DNA. This process takes place during the cycle sequencing reaction, a process that each sample we receive must undergo in order to become amplified and fluorescently labeled for detection on our sequencers.

In this cycle sequencing reaction, template and primer are combined together with a reaction mixture composed of dNTPs, fluorescently labeled ddNTPs, Amplitaq FS polymerase enzyme and buffer. The cycle sequencing reaction is composed of three steps - denaturation, annealing and extension - and takes place in a thermal cycler, an instrument that allows for controlled heating and cooling of our reactions. These steps are repeated for 35 cycles to ensure sufficient amplification of the labeled DNA, and takes about 3 1/2 hours to complete.

During the denaturation step, which occurs at 96ºC, the double-stranded template DNA is first separated into single-stranded molecules. At the annealing stage, the temperature is lowered to 50ºC so that the small primer molecules can find their complementary regions on the now single-stranded template DNA and hybridize, or anneal, correctly. The temperature is then raised to 60ºC (extension step) to allow the Taq polymerase enzyme to begin incorporation of nucleotides into growing chains of newly created fragments that are complementary to the single-stranded template DNA. These extension products begin at the end of the primer and extend in the 3’ direction. Chain termination occurs during this extension step. Our reaction mixture contains a mixture of deoxynucleotides (dNTPs) and dideoxynucleotides (ddNTPs), at concentrations that create a statistical probability that a dideoxynucleotide will be incorporated instead of a deoxynucleotide at each nucleotide position in the newly generated fragments. When a dNTP is incorporated, the new fragment will continue to grow. A ddNTP contains a hydrogen atom instead of a hydroxyl group at its 3’ end and cannot participate in further extension. Therefore, when a ddNTP is incorporated, further chain elongation is blocked and this results in a population of truncated products of varying lengths.

When separated by electrophoresis, a "ladder" of these truncated products will form, each differing in size by one nucleotide, with the smallest terminated fragments running fastest on the gel. There are four different ddNTPs that correspond to each of the four DNA nucleotides, and each ddNTP has a different color. Thus, each truncated fragment will contain a fluorescently labeled ddNTP at its 3’ end, and the sequencing ladder will be composed of colored bands. The sequence can then be determined by correlating the color of a band on the gel with its specific ddNTP, and the order in which they ran on the gel. So, the first and smallest band visualized will correspond to the first labeled nucleotide incorporated immediately adjacent to the primer. The second band will be the fragments that consist of 1 unlabeled dNTP and 1 labeled ddNTP that terminated those particular growing chains. The third band will be made up of 2 unlabeled dNTPS followed by 1 labeled ddNTP, and so on up the gel.

Once the sample has been amplified and labeled, it must be electrophoresed for separation of the labeled fragments and their visualization. As mentioned before, the ddNTPs are fluorescently labeled. The attached dyes are energy transfer dyes and consist of a fluorescein energy donor dye linked to an energy acceptor dichlororhodamine dye. This energy transfer system is much more sensitive than a single dye system and allows us to use less DNA for detection and, in addition, allows us to now sequence very large DNA molecules, such as BACs, PACs and even some bacterial genomic DNA, that previously less sensitive methods were unable to manage.

These dye-labeled fragments are loaded onto the sequencers and during electrophoresis, migrate either through the polyacrylamide gel or liquid polymer and are separated based on their size. Towards the end of the gel or the capillary, they pass through a region that contains a read window, behind which a laser beam passes back and forth behind the migrating samples. This laser excites the fluorescent dyes attached to the fragments and they then emit light at a wavelength specific for each dye. This emitted light is separated according to wavelength by a spectrograph onto a cooled, charge-coupled device, or CCD camera, so that all four fluorescent emissions can be detected by one laser pass. The data collection software collects these light intensities from the CCD camera at particular wavelength bands, or virtual filters, and stores them onto the sequencer’s computer as digital signals for processing. The analysis software then interprets the fluorescent intensity at each data point and assigns its base call interpretations.

Instrumentation in our laboratory

The Roswell Park DNA sequencing laboratory currently utilizes two ABI PRISM 3130XL Genetic Analyzers. They are automated capillary DNA sequencers manufactured by Applied Biosystems. Each 3130 uses a 16-capillary array, which can run 176 samples in 24 hours using our current configuration or up to 384 samples in 24 hours with a protocol which gives shorter read lengths, but allows for maximum throughput.

With the 3130, sample loading is based on electrokinetic injection. When a plate of samples is placed onto the instrument’s loading deck, the 16-capillary array dips into the first sixteen sample wells, voltage is applied and negatively charged ions, which are primarily the DNA extension products but which may also include interfering substances such as salts, are injected into the capillary array. This is one reason why DNA must be especially clean for runs on the 3130 as smaller negatively charged ions, such as salts, are preferentially injected and can interfere with the migration of the DNA fragments.

A liquid flowing polymer passes through the 16 capillaries in the array to act as the separation matrix. A fresh aliquot of polymer is pushed into the capillaries before each run, flushing out the polymer from the previous run. Previously, our laboratory used the longest capillary array provided for the 3100, 80 cm in length, and the Applied Biosystems POP-4 polymer for the separating medium. This combination gave around 900 bases of useful sequence, and data for sixteen samples could be acquired in 3 hours, 40 minutes with this array and polymer. However, we have since developed modified run protocols that utilize Applied Biosystems newest polymer, POP-7, and a 50 cm array. This polymer was developed specifically for ABI’s high-throughput sequencer (the model 3730) and has been shown to provide longer read-lengths coupled with shorter run times but was not supported for use on the 3100. Prior to upgrading to the 3130 models, we were able to obtain, on average, over 900-950 Phred Q20 bases with our “long-read” protocol and 850-900 Q20 bases with our “standard-read” protocol using our modified protocols. The “long-read” protocol has a run time of 2 hours, 45 minutes. Our “standard-read” protocol is complete in 2 hours, 5 minutes. Some of this work has been presented in Biotechniques. (2004 Jun36(6):932-3). With the 3130 upgrades, we are able to get similar read lengths and quality scores in a somewhat shorter period of time.

Electrophoresis is facilitated by a high-voltage electrical circuit in the 3130. The charge is conducted through the circuit by DNA and ions in the polymer, ions in the buffer used in the instrument, and through electrical wires and electrodes. The 3130 can evenly and efficiently dissipate heat from this voltage because of the large surface area of the fused silica capillaries. It is this attribute that allows for high voltage to be applied during electrophoresis, and as a result, contributes to significantly decreased run times without loss of resolution. Older sequencers such as the Applied Biosystems 377, which employed a slab gel in order to separate DNA fragments, could not dissipate heat as well and had to be run considerably slower and longer to attain read lengths similar to those on the 3130.

Sequencing fragments pass through a detection cell at the end of the array. Shorter fragments migrate more quickly than longer fragments and pass through the detection cell in this order. An argon-ion laser beam causes the electrons on the dyes attached to the DNA fragments to be excited to a higher energy state. When they return to their ground state, they fluoresce. The emitted fluorescence is captured by a charge-coupled device (CCD) camera. This camera is composed of a silica chip with thousands of pixels which store an electrical charge proportional to the intensity of the fluorescence that reaches it. The CCD camera then converts the fluorescence information into electronic data which is transferred to a computer for processing. Software processes this data in order to create an electropherogram where each peak represents one fragment of DNA sequencing product.

What kinds of DNA can we sequence and how much do we need?

The DNA sequencing laboratory can sequence a variety of DNA samples, including plasmids, cosmids, PCR products, single-stranded phage DNA, and BAC or PAC clones. When submitting DNA for sequencing, and if you are providing your own sequencing primers, concentrations for the various types of DNA should adjusted as follows:

DNA type Concentration Required Total used in a reaction
Plasmid 200-250 ng/ul 500-700 ng
Cosmid 200-250 ng/ul 500-700 ng
PCR Product 10-20 ng/ul 10 ng/100 bp
Single-Stranded Phage 100-150 ng/ul 300 ng
BAC/PAC 300-400 ng/ul 700-900 ng
Your Custom Primer 1 uM = 1 pmol/uL =

* Please adjust your DNA and primer concentrations accordingly to meet our concentration requirements.

At the concentrations listed above, we will use approximately 3-4 ul per reaction. Also, these measurements do not need to be exactly 200ng/ul or 10 ng/ul or whichever is applicable - these are approximations of how much we’d like and if you are off by a little - say, give or take 30 ng/ul for plasmids, 2-3 ng/ul for PCR products- it will still probably be fine as there is a range of concentrations we can use and still acquire good sequence data. And if your yields are much lower than our requirements as you do have a couple of options. First, if you have access to a vacuum centrifuge, aliquot out a volume that contains the TOTAL amount that we would use, dry it down and then resuspend in water to the concentration we would like. For example, if your initial plasmid DNA concentration is only 50 ng/ul, take out 10-15 ul, dry it down and then add 3 ul of water. If you don’t have access to a vacuum centrifuge, then make a note in the comments field of your request form and we will use a greater volume of your sample in our cycle sequencing reaction.

However, keep in mind that the maximum volume of DNA we can add to a sequencing reaction is 10 ul, so in order to attain our minimum total amount of DNA your concentration cannot be much less than 50 ng/ul for plasmids or cosmids, 70-90 ng/ul for BAC and PAC samples. If your sample concentration is lower than that, and you don’t have access to a vacuum centrifuge, then let us know and we can dry it down for you in our centrifuge, or we will show you how to do it. Alternatively, you can do an ethanol precipitation and resuspend in a smaller volume to concentrate your samples but remember to remove all traces of ethanol.

Generally, for PCR products, product yield isn’t a problem if your PCR conditions are optimized and robust. However, if you have a large PCR product that needs to be sequenced, that will require more DNA (for example, a 2 kb PCR product would require 100-150 ng of DNA), so you may want to scale up your reaction or pool multiple reactions together. In addition, when providing your own custom primers, please adjust their concentration to 1 uM, or 10 uM for BAC/PAC samples. At these concentrations, we will use 4 ul of primer per reaction. We provide a number of the more common vector primers, free of charge, for sequencing. Please check the link at Primers we provide to see the complete list. And whenever possible, please provide enough DNA and primer for two reactions in case we need to repeat a reaction for any reason.

Choosing your host strain

Choosing your host strain for your template preparation is an important consideration, as some strains will yield significantly better quality DNA for automated sequencing. Certain E.Coli strains may release certain factors, such as endonucleases and large amounts of carbohydrates, during their lysis that can be inhibitory to DNA preparation and sequencing. In addition, certain cellular components, such as positively charged polysaccharides, can compete with the DNA for binding in silica-based DNA preparation methods that are commonly used in many commercial kits. It is always recommended that you check with the manufacturers of specific strains to determine if there are any known difficulties that might arise when using their strains with your applications. Host strains that generally give reliable data include DH5alpha, DH10B, DH1, C600, XL1-Blue, XL10-Gold, TOP-10 and NM294. XL1 Blue grows slower than most strains and can lead to decreased DNA yield, but usually still works well. Strains such as JM101, JM83, HB101, TB1 and TG1 are NOT recommended as they can release large amount of inhibitory factors during lysis that can lead to poor quality DNA that will require extra purification steps.

Other factors to consider when preparing to grow up cultures are media choice and antibiotic selection. Overgrowth of cells should be avoided as cells begin to lyse fairly rapidly after reaching stationary phase, releasing large amounts of degraded chromosomal and plasmid DNA as well as polysaccharides that are difficult to separate away from plasmid DNA. Standard LB media or buffered Richer media is usually recommended, as an overnight culture of 10-12 hours will generally give an appropriate cell density for loading onto columns or membranes, and in addition will have a much lower proportion of degraded cellular materials and contaminating DNA. It’s best to avoid media such as TB or 2xYT as they will reach stationary phase much sooner, and thus an overnight culture will give an inappropriately high cell density as well as a greater percentage of inhibitory substances. If you should have to use media such as these, it’s advisable to monitor culture time and cell number to avoid DNA contamination and column overloading. One trick to try is washing your pelleted cells with 0.5M NaCl just prior to lysing to help remove polysaccharides and media materials that may co-elute with your DNA. Resuspend your pellet in the 0.5 NaCl solution, vortex well and spin again, removing the supernatant when finished.

Antibiotic selection should be maintained throughout to avoid growth of cells that do not contain plasmid. In the absence of effective selection, cells that do not contain plasmid will outgrow cells that do, leading to poor yield of the desired DNA. In addition, some plasmids will metabolize certain antibiotics, such as ampicillin, during cell growth so it is important to provide proper antibiotic concentrations to minimize depletion. Antibiotics should be aliquoted and frozen to maintain optimal efficacy.

Template preparation and purification

Template purity and concentration are the two most important factors in obtaining good quality sequence data. Template purity and concentration are the two most important factors in obtaining good quality sequence data. No, this is not a typo, it’s just that we can’t stress this principle enough when it comes to automated fluorescent sequencing, especially on our 3130s which require even more stringent cleanup methods. Fluorescent sequencing is very sensitive to certain contaminants in the DNA sample and DNA that is considered clean enough for many other molecular biological procedures such as PCR, cloning, or even manual radioactive sequencing, is NOT necessarily clean enough for fluorescent sequencing and can lead to poor quality data or sometimes no data at all. In this section, we list our recommendations on the best ways to clean up and quantitate your DNA, depending on the type of DNA you are trying to sequence.

Qiagen kits are one of the most universally accepted methods for giving consistently high quality DNA for automated sequencing. This is not to say that there are not other methods that do not work - we’ve listed a few other recommended protocols - but in our experience, Qiagen cleanup methods just have the best track record for reliability and consistency in their final product.

Another option that is available for DNA purification that gives template of high quality for automated DNA sequencing is the Mini-Plasmid DNA Isolation kit available from the Biomedical Research Service Center (BRSC) at SUNY Buffalo. This kit is "a modified and streamlined alkaline lysis mini-prep method from Sambrook et al. (Molecular Cloning, CSH, NY) that eliminates the need for additional RNase treatment and phenol-chloroform extraction. Plasmid DNA can be easily and rapidly purified from an overnight E. coli culture in 15 min with a typical yield of 1 – 5 mg DNA per ml of culture. The DNA is suitable for restriction digestion, probe preparation, and PCR analysis. DNA to be used for sequencing analysis is subjected to an additional 5-min step of PEG precipitation (PS-4) to remove any trace amount of RNA. High-quality DNA sequencing reads can be easily obtained (almost 1,000 bases resolved per sequencing run) if the plasmid DNA is isolated from a suitable E. coli host strain. The kit (sufficient for 300 mini-preps) is stable for at least one year if handled and stored properly." (taken from the BRSC website). At $95 for 300 plasmid minipreps, this kit is significantly cheaper than Qiagen kits (about one-fourth the cost), rapid and very simple to use. We have seen, first-hand, that the quality of DNA obtained from this kit is excellent for sequencing and we would recommend the use of this kit for DNA purification. The BSRC is a local SUNY resource that promotes collaborative translational research, develops and commercializes innovative products, and manufactures quality research reagents for the broad life science community. For more information about the BSRC and it's products, you can check out their website at: http://www.acsu.buffalo.edu/

A link to the mini-plasmid DNA isolation kit protocol can be found at: Mini-plasmid prep protocol.

Plasmid templates

Considerations when cleaning up plasmid preps

Poor template quality is one of the most common reasons for bad sequence data, as mentioned above, and is a prime consideration when choosing a plasmid cleanup method to give DNA of optimal purity for automated sequencing. Plasmid template quality can be affected by a variety of factors and contaminants including the following:

  • Salts or organics left over from template preparation
  • Presence of cellular components such as RNA, proteins, polysaccharides or chromosomal DNA
  • DNA that has degraded while in storage
  • Silica fines that carryover from template preparation kits that utilize loose resin or silica solutions

Here is a table of allowable contaminants and their acceptable concentration ranges that will still allow for good sequencing data:

Contaminant Amount Tolerated in Sequencing Reaction
RNA 1 ug
PEG 0.3%
NaOAc 5-10mM
Ethanol 1.25%
Phenol 0%
CsCl 5mM
EDTA 0.25mM

Some comments on the effects of some of these contaminants are listed below:

RNA - As you can see from the value in the chart above, a significant amount of RNA can actually be tolerated in the sequencing reaction. It is a common contaminant in plasmid minipreps, especially when columns are overloaded and the capacity of the lysis buffers is exceeded - overloading can be a problem with Qiagen minipreps. One of the ways RNA can potentially interfere is when quantitating your DNA preps by spectrophotometer- RNA also absorbs at 260 and when there is a large amount present, it can really throw off the accuracy of your concentration. It’s best, then, to treat your template preparations with RNase or high salt precipitation and also to quantitate your samples both by gel as well as spectroscopically.

PEG- the presence of residual polyethylene glycol in the template prep can have an inhibitory effect on the cycle sequencing Taq polymerase enzyme and lead to weak signal.

Salts- the processivity of the Taq polymerase used in the cycle sequencing reaction declines in the presence of high amounts of salts. Salt contamination in DNA preps may result from coprecipitation of salts in alcohol precipitations, insufficient removal of supernatant after precipitations or an incomplete wash of the pellet with 70% ethanol. Careful technique should be used when precipitating with alcohol. It has also been demonstrated that acetate ions, as opposed to sodium, potassium or chloride ions, are the most inhibitory in sequencing reactions. When using potassium acetate or sodium acetate, concentrations over 20 mM led to complete failure of the sequencing reactions, while concentrations of 60mM of sodium chloride were required before complete inhibition. Salts can be inhibitory when we sequence samples on our gel-based 377 but are even more problematic when running samples on our 3100 capillary system as these smaller, charged salts are preferentially injected and interfere with the migration of the DNA samples.

Ethanol- ethanol contamination can occur when the sample is insufficiently dried after precipitation or when carried over in an ethanol-containing wash buffer used in some DNA isolation procedures. Contamination with 10% or greater concentrations of ethanol usually leads to failure of the DNA sequencing reaction. Complete drying of the DNA samples is required to remove these traces of ethanol.

Phenol- phenol may be carried over from DNA alkaline lysis methods that utilize phenol and chloroform to remove proteins and other cellular contaminants from cell lysates. Phenol cannot be tolerated in the cycle sequencing reaction as it denatures proteins and will thus degrade the Taq polymerase enzyme used in the cycle sequencing reaction. Chloroform does not have the strong denaturing properties of phenol and doesn’t appear to adversely affect the sequencing reaction.

Cesium chloride - When using a cesium chloride ultracentrifugation density gradient protocol, one can obtain DNA of very high quality suitable for automated sequencing. HOWEVER, it is strongly recommended that you either perform dialysis followed by ethanol precipitation (best method) or do a minimal room temperature isopropanol precipitation to remove all traces of residual cesium chloride as the cesium can inhibit the Taq polymerase used in the cycle sequencing reaction.

EDTA - EDTA can chelate the magnesium required by the Taq polymerase in the cycle sequencing reaction, so when submitting samples, it is best to always have them diluted or resuspended in sterile ddH20 or 1X Tris buffer. Suspension in TE buffer is not recommended, though people have done it and many times there is not a problem. However, providing template DNA in water is an easy thing to do and if there is a problem with your sequence quality, the fact that there is no EDTA in your sample is one potential problem we can eliminate right away.

Qiagen kit principles

As mentioned above, Qiagen makes kits for extracting and purifying all types of DNA, including plasmid DNA. The Qiagen plasmid miniprep kits contain an anion exchange resin that consists of positively charged DEAE groups that preferentially bind the negatively charged DNA backbone. Once the DNA is bound to the resin, other contaminants such as salts, proteins, RNA and carbohydrates are washed away in a low-salt buffer. The DNA is eluted in a second step, using a high-salt buffer to dissociate it away from the resin. This kit is designed to give the highest purity DNA and is somewhat more costly. Another kit offered by Qiagen is the Qiaprep kit which uses a silica gel-based membrane to bind the DNA in the presence of high concentrations of chaotropic salts. Salts are removed with an ethanol wash and the DNA is then eluted with a low ionic strength Tris buffer. This method comes in a spin column format that can be used either in a centrifuge or on a vacuum manifold. This kit gives DNA of a purity that is suitable for sequencing and is also less expensive. However, there are some factors that one should keep in mind when using the Qiagen kits, or any other column-based miniprep kits. First, do not overload the columns - RNA and polysaccharides can compete with the DNA for binding on the column resin and lead to low yields of the desired plasmid DNA. Follow directions for cell growth and use LB media as recommended. Secondly, do not forget to wash the DNA with 70% ethanol following the isopropanol precipitation in order to remove excess salts. Washing two to four times with ethanol can improve DNA quality, and, in addition, extra washes with the binding solution can help remove other contaminants. Thirdly, warming your elution buffer can help improve yield. And lastly, remove all traces of ethanol before resuspending the final product in water.

Alkaline lysis/PEG methods

Bacteria can be lysed to release plasmid DNA by a number of methods including treatment with detergents, organic solvents, heat or alkali. When attempting to isolate large plasmids (>15kb) that are more susceptible to damage during the lysis procedure, a gentler method, such as equilibrium centrifugation or lysis involving a detergent like SDS, should be used. When lysing cells to harvest smaller plasmids, stronger methods, such as alkali treatment, can be utilized. These treatments cause the linear chromosomal DNA of the host to denature and degrade. Closed circular plasmid DNA strands do not separate completely as they are topologically intertwined and when conditions are returned to normal, the strands reanneal precisely in native conformation as superhelical molecules.

Standard alkaline lysis protocols, when coupled with PEG precipitation, can lead to small plasmid DNA of sufficient purity for use in automated fluorescent sequencing. The following is a detailed protocol for alkaline lysis/PEG treatment that is recommended by Applied Biosystems, manufacturers of our DNA sequencers.

Note: To minimize shearing of contaminating chromosomal DNA, do not use a vortex during this procedure.

  1. Pellet 1.5 -4.5 ml aliquots of culture for 1 minute in a microcentrifuge at maximum speed.
  2. Remove the supernatant by aspiration.
  3. Resuspend the bacterial pellet in 200 ul of GET buffer by pipetting up and down.
  4. Add 300 ul of freshly prepared 0.2N NaOH/1% SDS. Mix the contents of the tube by inversion. Incubate on ice for 5 minutes.
  5. Neutralize the solution by adding 300 ul of 3.0M potassium acetate, pH 4.8. Mix by inverting the tube. Incubate on ice for 5 minutes.
  6. Remove cellular debris by spinning in a microcentrifuge at maximum speed for 10 minutes at room temperature. Transfer the supernatant to a clean tube.
  7. Add RNase A (DNase free) to a final concentration of 20 ug/ml. Incubate the tube at 37C for 20 minutes.
  8. Extract the supernatant twice with chloroform:
    1. add 400 ul of chloroform
    2. mix the layers by inversion for 30 seconds
    3. centrifuge the tube for 1 minute to separate the phases
    4. transfer the upper aqueous phase to a clean tube.

    Cesium chloride purification

    Purification of plasmid DNA using a density gradient method such as equilibrium centrifugation in CsCl-ethidium bromide gradients can yield ultrapure DNA and is especially useful for purifying large plasmids or small plasmids that are to be used in more rigorous experiments, such as biophysical measurements. This protocol takes advantage of the fact that ethidium bromide will intercalate in different amounts when it encounters linear or closed circular DNA molecules. Due to the differential binding of this dye, the buoyant densities of linear and closed circular DNA that are formed after centrifugation will be different in CsCl gradients containing saturating amounts of ethidium bromide, thus allowing effective separation of plasmid DNA away from contaminating chromosomal DNA, nicked circular plasmid DNA, RNA and proteins. This method has long been the gold standard for obtaining very pure DNA, but it is rather time-consuming and expensive. Commercial kits or modified alkaline lysis methods provide DNA of adequate quality for automated sequencing, and so eliminate the need for such a stringent method. However, if cesium chloride purification is your method of choice, protocols can be found in the Maniatis Molecular Cloning manuals.

    Some final notes on plasmid purifications

    Boiling methods of DNA extraction are not acceptable for automated sequencing unless they are phenol/chloroform extracted to remove all proteins. Endonucleases are not always completely inactivated during the boiling procedure and plasmid DNA can be degraded in the presence of Mg++ in subsequent incubation procedures.

    And if, in the end, your plasmid prep is still not of optimal purity for good sequence data from our sequencers, here are some recommendations for further cleanup steps that may help:

    • Purify your DNA by ultrafiltration, using the Centricon-100 Micro-concentrator columns. See protocol at Ultrafiltration/Molecular weight cutoff columns.
    • Purify by further extraction:
      1. Extract the DNA twice with 1 volume of chloroform or chloroform:isoamyl alcohol (24:1 v/v)
      2. Add 0.16 volumes of 5M NaCl and 1 total volume of 13% PEG.
      3. Incubate on ice for 20 minutes, then centrifuge at maximum speed in a microcentrifuge at 2-6C for 20 minutes.
      4. Rinse the pellet twice with 70% ethanol.
      5. Dry the pellet in a vacuum centrifuge for 3-5 minutes or to dryness.
    • Purify using minimal isopropanol precipitations:
      1. Add 0.5 volume of 7.5M ammonium acetate or 0.1 volume of 3M sodium acetate to a dilute DNA solution (<100 ng/ul) and mix well.
      2. To this total volume, add 0.6 volume of room temperature isopropanol and mix well.
      3. Incubate at room temperature for 5 minutes, then centrifuge at maximum speed to pellet the DNA.
      4. Fill tube with 70% ethanol and do a complete wash of the pellet.
      5. Dry the pellet in a vacuum centrifuge for 3-5 minutes or to dryness.

    PCR products

    Considerations when cleaning up PCR products

    When submitting PCR products for direct sequencing, it is essential to separate the PCR product away from potentially interfering substances that may remain in solution after the PCR reaction. These contaminants can sometimes participate, and thus interfere, in the cycle sequencing reaction and lead to poor quality data or no data at all. Most importantly, primers and dNTPS should be removed. If both forward and reverse PCR primers remain in solution, they will both act as sequencing primers, resulting in multiple peaks from beginning to end as each primer can anneal to complementary strands with different nucleotide composition, and thus lead to overlapping fragments and unreadable data. Excess dNTPs should also be removed as they will upset the specific ratios of dNTPs/ddNTPs required for optimal extension and termination in the cycle sequencing reaction.

    Another consideration when choosing your PCR cleanup method is the determination of the presence of a single band or multiple bands that may result from your PCR reaction. Once the PCR reaction is completed, you should run a small aliquot on an agarose gel to assess its quality. If you have a single specific band, then it will be sufficient to remove excess PCR reactants using one of the methods listed below. If your PCR reaction generates multiple bands, an excessive amount of primer-dimers, or low-intensity smearing, it will then be necessary to run your PCR product out on a gel, excise your band of interest, and purify it away from the gel. Alternatively, you may want to spend some time optimizing your PCR reaction to eliminate the presence of these additional bands or artifacts and thus save yourself some downstream time in gel purifying your PCR products for sequencing.

    Regardless of which purification method you choose, you should always quantitate your PCR product after purification as no cleanup method will give 100% recovery. Any quantitative estimates you make when initially running your PCR product out on an analytical gel will be inaccurate as you will inevitably lose some product during purification, especially when gel purifying.

    Single PCR band

    Qiaquick PCR purification kit

    Qiagen makes PCR purification kits that come in either a microcentrifuge or vacuum manifold format, and can process anywhere from one PCR product using spin columns up to 96 samples in a 96 well plate. Fragments ranging in size from 100 bp up to 10 kb can be purified away from primers, dNTPs and salts that can interfere in the sequencing reaction. The protocol is simple and involves binding of DNA to the column or well membrane, washing away of contaminants, and elution of the DNA from the membrane. One nice thing about this kit is there are no extra steps to remove any mineral oil that may be present in the PCR solution.

    Exonuclease I/Shrimp Alkaline Phosphatase

    ExoSAP-IT is a product manufactured by USB Corporation and is an excellent method for purifying PCR reactions that produce one specific band. The Exonuclease I component functions to hydrolyze any residual single-stranded primers and any single-stranded DNA fragments produced in the PCR reaction. The Shrimp Alkaline Phosphatase degrades any excess dNTPS remaining in the reaction. The ExoSAP mixture is added directly to the PCR reaction, incubated at 37ºC for 15 minutes for hydrolysis of single-stranded DNA and dNTPS, and then incubated at 80ºC for 15 minutes to inactivate the enzymes.

    Ultrafiltration/Molecular weight cutoff columns

    Amicon Microcon and Centricon devices, which can be purchased from Millipore, are available with 30,000 and 100,000 molecular weight cut-off [MWCO] membranes and can separate primers from the synthesized PCR product. When purifying smaller PCR products (170 bp or less), choose the Centricon 30. For larger PCR products, the Centricon 100 works well. They will concentrate and desalt the DNA simultaneously, typically removing over 90% of the non-incorporated primers and dNTPs. When using either device, a small sample cup, or reservoir, is fitted into a collection tube. The reservoir contains the size-exclusion membrane at its bottom surface. The PCR product is loaded into the sample reservoir along with a suitable volume of distilled water (400 ul when using the Microcon filter, 2 ml for the Centricon), and then spun in a centrifuge. Salts, dNTPs and primers are spun through into the collection tube. To recover purified DNA, remove the sample reservoir from the vial, invert into a new tube and spin once again. The clean PCR product will then be captured in the second collection vial. Due to the increased wash volume and concentration factor, one spin in a Centricon device removes >95% of a 25 bp primer. A second spin removes up to an additional 4%. The Microcon filter will benefit from additional washes - one wash removes approximately 87% of primers, while three washes will filter away between 95%-99% of standard PCR-size primers.

    Ethanol precipitation

    If performed carefully, ethanol precipitation will yield PCR products of adequate purity for direct sequencing. However, if not done properly, residual primers and dNTPS, as well as any remaining ethanol, will give poor sequence data so be sure to remove supernatant carefully and completely, and keep your eye on the pellet.

    Both protocols listed below are for 50 ul PCR reactions, which tend to give better yields after purification, especially for larger products. For PCR reactions greater than 50 ul, proportionally scale up the reagents listed. For reactions less than 50 ul, add water to bring the volume to 50 ul. If mineral oil is present in the PCR reaction, add 100 ul of chloroform, (do not mix with PCR solution), spin at 10,000 RPM for 30 seconds and then aspirate supernatant into a clean tube. The chloroform/mineral oil component will settle to the bottom.

    • Protocol 1
      Thoroughly mix 25ul of 7.5M ammonium acetate with 190 ul 95% ethanol. Add purified supernatant (if mineral oil was previously used) or your PCR solution to this mixture. Vortex and place on ice, or at -20ºC, for 30 minutes to precipitate PCR products. Centrifuge at 10,000 RPM for 10 minutes at 4ºC. Completely remove supernatant, and dry the pellet, either by air-drying or in a vacuum centrifuge (do not overdry, a few minutes in a vacuum centrifuge will be fine). Resuspend in 20 ul of water.
    • Protocol 2
      Thoroughly mix 5 uL of 3M sodium acetate, pH 4.6, with 100 uL of 95% ethanol. Add purified supernatant or your PCR solution to this mixture. Vortex and place on ice, or at -20ºC, for 30 minutes to precipitate PCR products. Spin tubes in a microcentrifuge at maximum speed for 20 minutes. Completely remove supernatant and discard. Add 300 ul of 70% ethanol to rinse the pellet, vortex briefly, and spin for an additional 5 minutes at maximum speed. Completely remove supernatant and discard, being careful not to lose the pellet. Dry the pellet and resuspend in 50 ul water.

    Multiple PCR bands

    Gel purification

    When purifying a PCR product by gel filtration, either standard or low melting point agarose with TBE or TAE buffer may be used. However, it is important to use a high quality agarose such as FMC SeaPlaque or SeaPrep to minimize carryover of unwanted contaminants that may inhibit the sequencing reaction. When purifying the PCR band, the gel containing the PCR product of interest is first visualized under a UV light source and the band is then excised out of the agarose gel with a clean, sharp razor - minimize the amount of agarose that is cut out with the band, even if it means losing a bit of DNA. Tiny bits of agarose can have an adverse affect in the sequencing reaction. In addition, it is important to remember that UV light is damaging to your DNA - damage can occur in the time it takes you to cut out your band and it’s effect can become apparent when sequencing your DNA as you may see dramatically shortened read lengths. So whenever possible, reduce the intensity of the UV light source, reduce your exposure times or use a 366 nm UV source if available. Placing a glass plate between the light source and your gel can also help lessen the degrading effect of the UV light on your DNA. Once your band of interest has been isolated from the gel, numerous methods can be used to separate the PCR product away from the agarose slice, including the Qiaquick Gel Purification kit (MOST HIGHLY recommended), Millipore Ultrafree kit with modified TAE buffer, Promega PCR Wizard preps and Bio101 Geneclean kits. Follow the manufacturer’s direction carefully.

    PCR products should be examined and quantitated on an agarose gel AFTER gel purification as well, as you will not get 100% recovery and your assumed PCR concentration for sequencing will not be accurate.

    BAC/PAC DNA

    Considerations when cleaning up BAC/PAC DNA

    When attempting to sequence large DNA templates such as bacterial artificial chromosomes (BACs) or P1 derived artificial chromosomes (PACs), the quality of DNA template becomes even more crucial. As with plasmid DNA purification, Qiagen kits give excellent results for purification of BAC DNA for sequencing and they provide a few different methods for cleaning up the template DNA, including Qiafilter cartridges and a Large-Construct kit, the latter being somewhat more involved. One modified protocol of the Qiafilter Plasmid Midi Kit, which has consistently given good quality BAC DNA for sequencing, has been kindly provided to us from Wenjie Wu and we will post his modifications here.

    • use 100 ml of BAC culture
    • P1 buffer: add 10 ml
    • P2 buffer: add 10 ml
    • P3 buffer: add 10 ml
    • after addition of P3 buffer, incubate the sample on ice for 15 min, then centrifuge at 4000 rpm for 30 minutes at 4ºC (no incubation at room temperature).
    • add the supernatant from the centrifuged sample to the cartridge and filter the samples into a clean tube
    • apply the filtered supernatant to the Qiagen-tip for further purification and follow remaining Qiagen protocol.
    • when eluting the DNA, first warm the QF solution to 65ºC and elute 5 x 1ml to maximize yield.

    Other BAC purification methods can be used, including CsCl purification, and some other protocols are available at the following websites:

    University of Oklahoma Advanced Center for Genome Technology:
    http://www.genome.ou.edu/DblAcetateProcV3.html

    Note: BAC DNA prepped on the Autogen instrument generally does not work well for sequencing

    Methods for Quantitation

    There are three methods commonly used for quantitating DNA - spectrophotometry, agarose gel analysis and fluorometry. These methods can also give you a sense of how clean your DNA is, although none of these methods will detect everything. When using the spectrophotometer, you will typically take readings at both A260 and A280, as DNA will absorb at A260 while protein absorbs at A280. These readings will give you an A260/A280 ratio. A good DNA A260/A280 ratio should be between 1.7-1.9. If you find your ratio is smaller, this could indicate contamination of your sample with proteins or organic chemicals. If possible, use a spectrophotometer that is capable of performing a wavelength scan from 220 nm to 330 nm as a scan may be useful in revealing other contaminants such as phenol, which is detectable at 230nm, and particulates in solution that may absorb at 325nm. When calculating your DNA concentration with a spectrophotometer, you will need to know that one A260 O.D. unit (absorbance) of double-stranded DNA contains 50 ng/ul. One O.D. unit is the amount of a substance dissolved in 1 ml that gives an absorbance reading of 1.0 in a spectrophotometer with a 1-cm path length. So to calculate the concentration of DNA you have you will multiply the absorbance value times the conversion factor of 50ng/ul times any dilution factor you may have used and this will give you the concentration of your sample in ng/ul or mg/ml. However, quantitation of your DNA by spec is probably the least accurate of the three methods as both RNA and contaminating DNA will also absorb at 260nm and can potentially lead to inaccurate values. Also, readings greater than 1.0 OD or less than 0.05 are probably not very accurate, limiting the sensitivity and range of this method.

    Quantitating by agarose gel electrophoresis tends to be more accurate as you can visualize any contaminating DNA or RNA. Purified DNA should run as a single band on an agarose gel, while uncut plasmid DNA will show three bands: supercoiled, nicked and linear. This method is also useful when quantitating small amounts of DNA such as PCR products. The DNA sample is run on a gel next to a mass ladder composed of fragments of known and varying concentrations and the intensity of your DNA band is visually compared to the intensities of the bands from the mass ladder to estimate DNA concentration. When using an agarose gel, you can’t detect the presence of proteins or chemicals. To get the most information about the amount and quality of your desired DNA, both spectroscopic and gel quantitation methods should be used together.

    Perhaps the most accurate method of measurement of the three is the use of a fluorometer. Dyes may be added to the DNA such as Hoechst 33258 Dye or Picogreen which intercalate into AT rich regions of double stranded DNA and are thus quite specific to double stranded DNA, avoiding the problems of contaminating RNA seen when using a spectrophotometer. Common contaminants present in DNA preparations such as salts, urea, ethanol, chloroform, detergents, proteins and agarose do not have significant affect on these assays and thus lead to a much more sensitive and precise method for quanititation. Fluorometric measurements can detect DNA levels as low as 5 ng/ul.

    Primer design

    There are many programs on the Internet that can help you choose primers, determine potential primer-dimer formation or hairpin structures, and calculate Tm. These programs can be especially helpful when you need to design multiple sets of primers to ensure coverage of a large region of DNA. Look at our section Useful Links for some primer design programs that we have found helpful. That being said, here are some common rules for sequencing primer design:

    • primer should be 18-24 nucleotides in length - this helps ensure good and specific hybridization to the template DNA. Long primers (>35-40bp) tend not to work quite as well in sequencing due to the increased chances of secondary structure formation
    • G/C content should be approximately 50%, but the range can be 30-80%
    • Tm should be between 50º-60ºC - if your Tm is much below that it may not hybridize properly, as the annealing temperature in our cycle sequencing reaction is 50ºC. If necessary, add a few more bases to your primer to increase its Tm. To calculate the Tm of your primer use the following formula:
      • Tm = 4(G + C) + 2(A + T)º C

      As a last note on ordering primers - while purity of a primer is important, it is generally not necessary to have ultrapure primers, such as those purified by HPLC or OPC cartridge, though that is, of course, optimal. As long as the oligo synthesis efficiency was high and the majority of your primer solution contains full-length primers, then simple desalting of your primers should be fine. Primers of poor synthesis quality will not work well for sequencing because, along with your full-length product, there will be a greater proportion of "failure" primers (i.e. truncated primers of less than full length, in varying sizes) that can also act as sequencing primers. Sequence data from these primers will be poor, ranging from unusable to minor "shadow" peaks under the proper peaks. See more in our Troubleshooting section to see how to recognize this problem. When submitting custom primers for sequencing, dilute them in water to the proper concentration.

      For the do-it-yourselfers

      For those selecting the economical choice of setting up your own sequencing reactions, certain things must be taken into consideration. First, you will need to purchase the kit to perform your cycle sequencing reactions. Cycle sequencing is the method required to amplify and fluorescently label your DNA so that it can be detected on our automated sequencers. Cycle sequencing is very similar to PCR, but with two major differences. In cycle sequencing, one primer is used instead of two, thus giving linear amplification of one labeled strand. In addition, a mixture of labeled and unlabeled dNTPs are used in the reaction mix to allow for fluorescent labeling of the DNA and fragment chain termination. Read our Principles of automated fluorescent DNA sequencing section for more detailed information on the basics of cycle sequencing. Once the reactions have been set up, you will need to run them in a thermal cycler to effect the incorporation of fluorescently labeled ddNTPs and amplification of your desired DNA. Once the cycle sequencing reaction is completed, you will need to purify the reaction to remove salts, unused reactants and, most importantly, unincorporated Dye Terminator molecules. If this step is not performed carefully, these excess dye molecules will migrate along with your DNA and cause what are known as “dye blobs” which can interfere with proper basecalling of your DNA. See out Troubleshooting section for an example of a dye blob.

      With that being said, we’ve provided below our recommendations and protocols, as well as a listing of some supplies and part numbers you’ll need to get started.

      Reagents

      Cycle sequencing reaction mixes must be purchased from Applied Biosystems, manufacturers of our sequencers. For best sequence quality and read length, we recommend the use of Big Dye Terminator chemistries, which is what we use in our sequencing reactions. Two standard chemistries are available for routine template sequencing, Big Dye Terminator v3.1 and 1.1. Part numbers are listed below:

      BigDye ® Terminator v3.1 Cycle Sequencing Kit
      (protocol ordered separately p/n 4337035)


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      Innovative technologies

      At Illumina, our goal is to apply innovative technologies to the analysis of genetic variation and function, making studies possible that were not even imaginable just a few years ago. It is mission critical for us to deliver innovative, flexible, and scalable solutions to meet the needs of our customers. As a global company that places high value on collaborative interactions, rapid delivery of solutions, and providing the highest level of quality, we strive to meet this challenge. Illumina innovative sequencing and array technologies are fueling groundbreaking advancements in life science research, translational and consumer genomics, and molecular diagnostics.

      For Research Use Only. Not for use in diagnostic procedures (except as specifically noted).


      Potential Beyond Drug Discovery

      Such next-generation sequencing approaches are used by pharmaceutical companies such as Novartis, as well as many academic centers, some of them involved in cancer genome sequencing consortiums, to generate lists of genes that are frequently mutated in certain cancer types, explains Barretina. “Once you are armed with that knowledge, you can start designing drugs that will actually target those genetic alterations in particular cancers.”

      As soon as a patient is diagnosed, we should quickly determine what’s driving the cancer and use that information to select an appropriate therapy.

      Wendy Winckler, Executive Director of Next-Generation Diagnostics at Novartis Institutes for BioMedical Research

      Cataloging genetic alterations of cancer cells was one of the principal goals behind the Cancer Cell Line Encyclopedia — a massive undertaking and collaborative project between Novartis and the Broad Institute, also based in Cambridge, Mass. The encyclopedia provides genetic information on approximately 1,000 different cancer cell lines originally derived from patients’ tumors. The data is released publicly so that researchers around the world can mine it. Cell lines are easy-to-use cancer models that allow researchers to investigate cancer biology and test the effectiveness of potential drugs against various types of tumors. The Cancer Cell Line Encyclopedia also contains corresponding data detailing which cell lines respond to which drugs.

      Giordano Caponigro, one of the leaders of the Cancer Cell Line Encyclopedia collaboration, is now senior director in Oncology Translational Research at Novartis. He explains how next-generation sequencing is increasing the value of this resource. “When the Cancer Cell Line Encyclopedia started, next-generation sequencing hadn’t really hit its stride, but as it went on we realized this was an important technology,” he says. “Shortly, whole-genome sequencing of the full set of cancer cell lines will be complete, which will help with the development of highly accurate diagnostic tools and effective new cancer treatments.”


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      V-pipe has a modular and extensible architecture. Users can design their own fully reproducible and transparent pipelines. Developers can test their own tools in a defined environment and contribute to the establishment of best practices for virus research and clinical diagnostics.

      V-pipe is freely available for download from GitHub. Further details on how to run the pipeline, as well as a test dataset, can be found on the dedicated Wiki pages of the repository, accessible through the V-pipe website’s Usage tab.

      Should you have any further question, please do not hesitate to contact us.


      New Approach to Nanopore Sequencing That Is Sure to CATCH Your Interest

      A group of scientists from Tel Aviv University recently performed an experiment utilizing Cas9-Assisted Targeting of CHromosome segments (CATCH) to isolate a gene typically associated with breast cancer, BRCA1, and used nanopore sequencing to map the gene.

      Experimental Fulcrum

      The team extracted 200kb strands of DNA from primary human peripheral blood cells—containing the 80kb BRCA1 gene and its adjacent sectors—and proceeded to process the strands with long-range amplification. Once the target sequence of 200kb was sufficiently enriched, the team began nanopore sequencing which allowed them to determine the order of nucleotides that comprised the section of interest, and even allowed for the registration of both coding and non-coding areas which may help shed light on less straightforward genetic determinants.

      Being able to extract, amplify, and analyze only 200kb rapidly expedites the ability to sequence a specific area within the human genome as compared to the massive 3 billion base pairs that make up all of our genetic information. “Due to high sequencing costs, cutting out only the region of interest from the genome focuses all your sequencing efforts and money toward this target without ‘wasting’ reads on DNA from the rest of the genome,” explains Yuval Ebenstein, Ph.D., a professor at Tel Aviv University.

      Techniques for mapping the genome are varied, including single-molecule real-time (SMRT) sequencing, nanopore sequencing, and, a little older but still relevant attributable to its higher accuracy, next-generation sequencing (NGS)—an umbrella term for technologies such as Illumina sequencing.

      However, while these methods have proven effective they are not without their limitations. NGS is hindered due to its inability to process structural aberrations greater in size than a few hundred base pairs. Long-read methods such as SMRT and nanopore sequencing are capable of detecting variations that NGS can’t, and both provide real-time viewing of data. Nanopore sequencing takes it a step further and is capable of analyzing native DNA without the need for PCR amplification techniques—which all other sequencing methods depend on. The elimination of prerequisite amplification is a huge advantage because it is often during PCR amplification that more errors and single nucleotide polymorphisms (SNPs) are produced. With that said, nanopore sequencing still has a ways to go in terms of achieving greater accuracy, while NGS has shown itself to be tried and true.

      Dr. Ebenstein points out that while nanopore sequencing is normally able to make use of native DNA, “CATCH isolates only a tiny fraction of the genome which means that very low amounts of DNA are recovered for library preparation. Currently, this implies that the DNA must either be isolated from a very large amount of cells or amplified to achieve the needed amount of material for sequencing.”


      CATCH nanopore sequencing promises to increase our genome mapping capabilities. [Tel Aviv University]

      Finance, Function, and Future

      What does all of this mean? NGS is incapable of processing reads bigger than a few hundred base pairs, while nanopore technology is capable of processing reads up to several kilo base pairs. However, an intrinsic obstacle to nanopore sequencing is that it can be expensive. It’s true that the upfront cost for Oxford Nanopore Technologies’ MinION retails for only $1,000, but while “the initial capital investment is indeed lower, the price per base on the MinION is still considerably higher than NGS,” notes Dr. Ebenstein, “although [nanopore sequencing] can obtain information that is inaccessible to NGS.”

      By utilizing CATCH, scientists can isolate and enrich smaller portions of the genome, as done in this experiment, by having the Cas9 cut at the sequence of interest’s flanking regions. It is possible to expedite the process, as nanopore sequencing can be used only on the cut-out region of interest rather than the entire genome. That was one of the crucial points that the team wanted to convey with this experiment.

      “CATCH brings nanopore sequencing closer to the clinic by offering targeted, cost-effective sequencing,” explains Dr. Ebenstein. “You only need sequence knowledge for two regions flanking your target, thus enabling the enrichment of large variable regions in the genome that are very complex and challenging for conventional approaches.”

      This is cause for excitement as CATCH furthers the potential of an already impressive technique for DNA sequencing. Nanopore sequencing is already highly valued for its ultra-long reads and real-time analysis capability, but with the incorporation of CATCH the entire process can occur even faster and with greater fiscal efficacy.

      “The long reads provided by nanopore sequencing allow for the characterization of structural variations in conjunction with smaller aberrations such as SNPs. Together, this information is critical for understanding the genetic causes of disease,” states Dr. Ebenstein.

      The results of the experiment were telling as to the capabilities and limitations of both NGS and nanopore sequencing. In their paper, published in Nucleic Acids Research as “Selective nanopore sequencing of human BRCA1 by Cas9-assisted targeting of chromosome segments (CATCH)”, the team shared data that “NGS discovered 177 SNPs. Nanopore data revealed 62% of the SNPs with only two SNPs not present at all in the nanopore data and the remaining 37% inconclusive.”

      “NGS has a lower error rate compared to nanopore. Therefore, it is more qualified to detect single base mutations,” says Dr. Ebenstein. “On the other hand, sufficient coverage (offered by CATCH) can overcome these errors by averaging over many reads. In addition, nanopore has better access to larger aberrations such as structural and copy number variants.”

      Yael Michaeli, Ph.D., School of Chemistry, Faculty of Exact Sciences at Tel Aviv University, mentions that their team “used a software that is more compatible with NGS reads and this in part may explain the inferior results of the nanopore.”

      It’s clear that there are still some areas of uncertainty surrounding nanopore sequencing and compared to the competition it still has to prove itself a little further in terms of reliability. Still, Dr. Ebenstein believes that “by now it seems quite clear that nanopore sequencing will find its place in routine genomic analysis pipelines. [However,] it is still more expensive than NGS, and should be chosen only when needed.”


      Next-Generation Sequencing (NGS)

      Next-generation sequencing (NGS) is a massively parallel sequencing technology that offers ultra-high throughput, scalability, and speed. The technology is used to determine the order of nucleotides in entire genomes or targeted regions of DNA or RNA. NGS has revolutionized the biological sciences, allowing labs to perform a wide variety of applications and study biological systems at a level never before possible.

      Today's complex genomics questions demand a depth of information beyond the capacity of traditional DNA sequencing technologies. NGS has filled that gap and become an everyday tool to address these questions.

      Next-Generation Sequencing for Beginners

      We'll guide you through the basics of NGS, with tutorials and tips for planning your first experiment.

      See What NGS Can Do For You

      NGS technology has fundamentally changed the kinds of questions scientists can ask and answer. Innovative sample preparation and data analysis options enable a broad range of applications. For example, NGS allows labs to:

      • Rapidly sequence whole genomes
      • Deeply sequence target regions
      • Utilize RNA sequencing (RNA-Seq) to discover novel RNA variants and splice sites, or quantify mRNAs for gene expression analysis
      • Analyze epigenetic factors such as genome-wide DNA methylation and DNA-protein interactions
      • Sequence cancer samples to study rare somatic variants, tumor subclones, and more
      • Study the human microbiome
      • Identify novel pathogens

      Accessible Whole-Genome Sequencing

      Using capillary electrophoresis-based Sanger sequencing, the Human Genome Project took over 10 years and cost nearly $3 billion.

      Next-generation sequencing, in contrast, makes large-scale whole-genome sequencing (WGS) accessible and practical for the average researcher. It enables scientists to analyze the entire human genome in a single sequencing experiment, or sequence thousands to tens of thousands of genomes in one year.

      NGS Data Analysis Tools

      Explore user-friendly tools designed to make data analysis accessible to any scientist, regardless of bioinformatics experience.

      Broad Dynamic Range for Expression Profiling

      NGS-based RNA-Seq is a powerful method that enables researchers to break through the inefficiency and expense of legacy technologies such as microarrays. Microarray gene expression measurement is limited by noise at the low end and signal saturation at the high end.

      In contrast, next-gen sequencing quantifies discrete, digital sequencing read counts, offering a broader dynamic range. 1,2,3

      Tunable Resolution for Targeted NGS

      Targeted sequencing allows you to sequence a subset of genes or specific genomic regions of interest, efficiently and cost-effectively focusing the power of NGS. NGS is highly scalable, allowing you to tune the level of resolution to meet experimental needs. Choose whether to do a shallow scan across multiple samples, or sequence at greater depth with fewer samples to find rare variants in a given region.

      NGS for COVID-19

      Next-generation sequencing is uniquely positioned in an infectious disease surveillance and outbreak model. Learn which NGS methods are recommended for detecting and characterizing SARS-CoV-2 and other respiratory pathogens, tracking transmission, studying co-infection, and investigating viral evolution.

      How Does Illumina NGS Work?

      Illumina sequencing utilizes a fundamentally different approach from the classic Sanger chain-termination method. It leverages sequencing by synthesis (SBS) technology – tracking the addition of labeled nucleotides as the DNA chain is copied – in a massively parallel fashion.

      Next-generation sequencing generates masses of DNA sequencing data, and is both less expensive and less time-consuming than traditional Sanger sequencing. 2 Illumina sequencing systems can deliver data output ranging from 300 kilobases up to multiple terabases in a single run, depending on instrument type and configuration.

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      In-Depth NGS Introduction

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      What Can You Do with Next-Generation Sequencing?

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      Genetics of COVID-19 Susceptibility

      This UK-wide study uses NGS to compare the genomes of severely and mildly ill COVID-19 patients, to help uncover genetic factors associated with susceptibility.

      Exploring the Tumor Microenvironment

      Researchers use single-cell techniques to study cancer microenvironments, to elucidate gene expression patterns and gain insights into drug resistance and metastasis.

      Using NGS to Study Rare Diseases

      Whole-exome and transcriptome sequencing prove beneficial in uncovering mutations and pathways associated with rare genetic diseases.

      Evolution of Illumina NGS

      Recent Illumina next-generation sequencing technology breakthroughs include:

        : The iSeq 100 System combines a complementary metal-oxide semiconductor (CMOS) chip with one-channel SBS to deliver high-accuracy data in a compact system. : This technology enables faster sequencing than the original 4-channel version of SBS technology, with the same high data accuracy. : This option offers an exceptional level of throughput for diverse sequencing applications. : Learn how the NovaSeq 6000 System offers tunable output of up to 6 Tb in

      History of Illumina Sequencing

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      Bring NGS to Your Lab

      The resources below offer valuable guidance to scientists who are considering purchasing a next-generation sequencing system.

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      Use our interactive tools to help you create a custom NGS protocol or select the right products and methods for your project.

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      References
      1. Wang Z, Gerstein M, Snyder M. RNA-Seq: a revolutionary tool for transcriptomics. Nat Rev Genet. 200910:57–63.
      2. Wilhelm BT, Landry JR. RNA-Seq—quantitative measurement of expression through massively parallel RNA sequencing. Methods. 200948:249–57.
      3. Zhao S, Fung-Leung WP, Bittner A, and Ngo K, Liu X. Comparison of RNA-Seq and microarray in transcriptome profiling of activated T cells. PLoS One. 2014169(1):e78644.

      Innovative technologies

      At Illumina, our goal is to apply innovative technologies to the analysis of genetic variation and function, making studies possible that were not even imaginable just a few years ago. It is mission critical for us to deliver innovative, flexible, and scalable solutions to meet the needs of our customers. As a global company that places high value on collaborative interactions, rapid delivery of solutions, and providing the highest level of quality, we strive to meet this challenge. Illumina innovative sequencing and array technologies are fueling groundbreaking advancements in life science research, translational and consumer genomics, and molecular diagnostics.

      For Research Use Only. Not for use in diagnostic procedures (except as specifically noted).


      Introduction

      Initial sequencing of the human genome took more than a decade and cost an estimated $70 million dollars (1). Sequencing for the Human Genome Project (HGP) relied primarily on automation of sequencing methods first introduced by Sanger in 1977 (2). Despite the successful use this technology to generate early maps of the human genome (3–5), the limitations of Sanger sequencing created a high demand for more robust sequencing technologies capable of generating large amounts of data, quickly, and at lower costs.

      Recognizing this need, the National Human Genome Research Institute (NHGRI) initiated a funding program in 2004 aimed at catalyzing sequencing technology development and with a goal of reducing the cost of genome sequencing to ∼$100,000 in 5 years and, ultimately, $1,000 in 10 years (6–8). The initiative has been widely successful to date, and a bevy of new technologies has emerged in the sequencing marketplace over the past 5 years. New technologies offer radically different approaches that enable the sequencing of large amounts of DNA in parallel and at substantially lower costs than conventional methods. The terms “next-generation sequencing” and “massive-parallel sequencing” have been used loosely to collectively refer to these new high-throughput technologies.


      CONCLUSION

      This study has proved that RNA ligases derived from T4-phage exhibit significant sequence specificity in their activity. The profiles of small RNAs are strongly dependent on the adapters used for sample preparation. In light of this, the current, popular, sRNA-seq protocols need revision. We find that a mix of adapters, with different sequence ends, permits a more accurate estimation of the amounts of individual miRNA sequences and their isoforms. The use of RNA ligases in other protocols, such as oligoribonucleotide circularization ( 38), should be reviewed for possible effects of the bias discussed in this study.


      Watch the video: Δημιουργία βάσης δεδομένων ΜySQL (January 2023).