There have been many changes in sequencing technology over the course of my scientific career. In one of the research labs I rotated in as a graduate student, I assisted a third-year grad student with a manual radioactive sequencing gel because, I was told, “every student should run at least one in their career”. My first job after graduate school was as a research assistant in a lab that sequenced bacterial genomes. While I was the one creating shotgun libraries for the DNA sequencing pipeline, the sequencing reaction was performed using dideoxynucleotides labeled with fluorescent dyes and amplified in thermal cyclers. The resulting fragments were separated by manual loading on tall slab polyacrylamide gels (Applied Biosystems ABI 377s) or, once the lab got them running, capillary electrophoresis of four 96-well plates at a time (ABI 3700s).
Sequencing throughput has only increased since I left the lab. This was accomplished by increasing well density in a plate and number of capillaries for use in capillary electrophoresis, but more importantly, with the advent of the short read, massively parallel next-generation sequencing method. The next-gen or NGS technique decreased the time needed to sequence because many sequences were determined at the same time, significantly accelerating sequencing capacity. Instruments have also decreased in size as well as the price per base pair, a measurement used when I was in the lab. The long-prophesized threshold of $1,000 per genome has arrived. And now, according to a recent tweet from a Nanopore conference, you can add a sequencing module to your mobile device:
Imagine you are traveling in your car and must pass through a mountain range to get to your destination. You’ve been following a set of directions when you realize you have a decision to make. Will you stay on your current route, which is many miles shorter but contains a long tunnel that cuts straight through the mountains and obstructs your view? Or will you switch to a longer, more scenic route that bypasses the tunnel ahead and gets you to your destination a bit later than you wanted?
Choosing which route to take illustrates a clear trade-off that has to be considered—which is more valuable, speed or understanding? Yes, the tunnel gets you from one place to another faster. But what are you missing as a result? Is it worth a little extra time to see the majestic landscape that you are passing through?
Considering this trade-off is especially critical for researchers working with human DNA purified from formalin-fixed paraffin-embedded (FFPE) or circulating cell-free DNA (ccfDNA) samples for next-generation sequencing (NGS). These sample types present a few challenges when performing NGS. FFPE samples are prone to degradation, while ccfDNA samples are susceptible to gDNA contamination, and both offer a very limited amount of starting material to work with.
One of the most critical parts of a Next Generation Sequencing (NGS) workflow is library preparation and nearly all NGS library preparation methods use some type of size-selective purification. This process involves removing unwanted fragment sizes that will interfere with downstream library preparation steps, sequencing or analysis.
Different applications may involve removing undesired enzymes and buffers or removal of nucleotides, primers and adapters for NGS library or PCR sample cleanup. In dual size selection methods, large and small DNA fragments are removed to ensure optimal library sizing prior to final sequencing. In all cases, accurate size selection is key to obtaining optimal downstream performance and NGS sequencing results.
Current methods and chemistries for the purposes listed above have been in use for several years; however, they are utilized at the cost of performance and ease-of-use. Many library preparation methods involve serial purifications which can result in a loss of DNA. Current methods can result in as much as 20-30% loss with each purification step. Ultimately this may necessitate greater starting material, which may not be possible with limited, precious samples, or the incorporation of more PCR cycles which can result in sequencing bias. Sample-to-sample reproducibility is a daily challenge that is also regularly cited as an area for improvement in size-selection.
Forensic analysts have long sought precision when determining time of death. While on crime scene investigation television shows, the presence of insects always seems to reveal when a person died, there are many elements to account for, and the probable date may still not be accurate. Insects arrive days after death if at all (e.g., if the body is found indoors or after burial), and the stage of insect activity is influenced by temperature, weather conditions, seasonal variation, geographic location and other factors. All this makes it difficult to estimate the postmortem interval (PMI) of a body discovered an unknown time after death. One way to make estimating PMI less subjective would be to have calibrated molecular markers that are easy to sample and are not altered by environmental variabilities.
Bacterial communities called microbiomes have been frequently in the news. The influence of these microbes encompass living creatures and the environment. Not surprisingly, research has focused on the influence of microbiomes on humans. For example, changes in gut microbiome seem to affect human health. Intriguingly, microbiomes may also be a key to determining time of death. The National Institute of Justice (NIJ) has funded several projects focused on the forensic applications of microbiomes. One focus involves the necrobiome, the community of organisms found on or around decomposing remains. These microbes could be used as an indicator of PMI when investigating human remains. Recent research published in PLOS ONE examined the bacterial communities found in human ears and noses after death and how they changed over time. The researchers were interested in developing an algorithm using the data they collected to estimate of time of death. Continue reading
Instruments can make our lives easier in the lab. Place your samples inside an instrument and let it do all the work—isolating nucleic acids or reading and analyzing a multiwell plate—while you walk away to read a new research paper or prepare for the next step in your experiment. However, with the array of machines now available to scientists worldwide, some confusion may result in the laboratory. Has this ever happened to you?
Copyright Ed Himelblau
Today’s blog is written by guest blogger, Douglas R. Storts, PhD, head of Research, Nucleic Acid Technologies, Promega Corporation.
Massively parallel sequencing (MPS), also called next generation sequencing (NGS), has the potential to alleviate some of the biggest challenges facing forensic laboratories, namely degraded DNA and samples containing DNA from multiple contributors. Unlike capillary electrophoresis, MPS genotyping methods do not require fluorescently-labeled oligonucleotides to distinguish amplification products of similar size. Furthermore, it is not necessary to design primers within a color channel to generate amplicons of different sizes to avoid allele overlap. Consequently, all the amplicons can be of a similar, small size (typically <275 base pairs). The small size of the amplicons is particularly advantageous when working with degraded DNA. Because the alleles are distinguished by the number of repeats and the DNA sequence, additional information can be derived from a sample. This can be especially important when genotyping mixtures. As previously demonstrated (1), this sequence variation can help distinguish stutter “peaks” from minor contributor alleles.
Because there is no reliance upon size and fluorescent label, significantly greater multiplexing is possible with MPS approaches. In addition to autosomal short tandem repeats (STRs), we can also sequence Y-STRs, single nucleotide polymorphisms (SNPs), and the mitochondrial DNA control region. The advantage to this approach is the forensic analyst does not need a priori knowledge whether a sample would benefit most from the different methods of genotyping.
Despite these major advantages, there are limitations to the near-term, broad deployment of current MPS technology into forensic laboratories. The limitations fall into four main categories: Workflow, costs, performance with forensically-relevant samples, and community guidelines. Continue reading
Real-Time (or quantitative, qPCR) monitors PCR amplification as it happens and allows you to measure starting material in your reaction.
This the last in a series of four blogs about Quantitation for NGS is written by guest blogger Adam Blatter, Product Specialist in Integrated Solutions at Promega.
When it comes to nucleic acid quantitation, real-time or quantitative (qPCR) is considered the gold standard because of its unmatched performance in senstivity, specificity and accuracy. qPCR relies on thermal cycling, consisting of repeated cycles of heating an cooling for DNA melting and enzyamtic replication. Detection instrumentation is capable of measuring the accumulation of DNA product after each round of amplification in real time.
Because PCR amplifies specific regions of DNA, the method is highly sensitive, specific to DNA, and it can determine whether a sample is truly able to be amplified. Degraded DNA or free nucleotides, which might otherwise skew your quantiation, will not contribute to the signal, and your measurement will be more accurate.
However, while qPCR does provide technical advantages, the method requires special instrumentation, specialized reagents and is a more time-consuming process. In addition, you will probably need to optimize your qPCR assay for each of your targets to achieve your desired results.
Because of the added complexity and cost, qPCR is a technique suited for post-library quantitation when you need to know the exact amount of amplifiable, adapter-ligated DNA. PCR is the only method capable of specifically targeting these library constructs over other DNA that may be present. This specificity is important because accurate normalization is especially critical for producing even coverage in multiplex experiments where equimolar amounts of several libraries are added to a pooled sample. This normalization process is essential if your are screening for rare variants that might be lost in background and go undetected if underrepresented in a mixed pool.
This is the third post in a series of blogs on quantitation for NGS applications written by guest blogger Adam Blatter, Product Specialist in Integrated Solutions at Promega.
Fluorescent dye-based quantitation uses specially designed DNA binding compounds that intercalate only with double stranded DNA molecules. When excited by a specific wavelength of light, only dye in the DNA-bound state will fluoresce. These aspects of the technique contribute to low background signal, and therefore the ability to accurately and specifically detect very low quantities of DNA in solution, even the nanogram quantities used in NGS applications.
For commercial NGS systems, such as the Nextera Rapid Capture Enrichment Protocol by Illumina, this specificity and sensitivity of quantitation are critical. The Nextera protocol is optimized for 50ng of total genomic DNA. A higher mass input of genomic DNA can result in incomplete tagmentation, and larger insert sizes, which can adversely affect enrichment. A lower mass input of genomic DNA or low-quality DNA can generate smaller than expected inserts, which can be lost during subsequent cleanup steps, giving lower diversity of inserts. Continue reading
For UV-Vis Spectrophotometry, light is split into its component wavelengths and directed through a solution. Molecules in the solution absorb specific wavelengths of light.
This is the second in a series of four blogs about Quantitation for NGS is written by guest blogger Adam Blatter, Product Specialist in Integrated Solutions at Promega.
Perhaps the most ubiquitous quantitation method is UV-spectrophotometry (also called absorbance spectroscopy). This technique takes advantage of the Beer-Lambert Law: an observation that many compounds absorb UV-Visible light at unique wavelengths, and that for a fixed path length the absorbance of a solution is directly proportional to the concentration of the absorbing species. DNA, for example has a peak absorbance at 260nm (A260nm).
This method is user friendly, quick and easy. But, it has significant limitations, especially when quantitating samples for NGS applications. Continue reading
This series of blogs about Quantitation for NGS is written by guest blogger Adam Blatter, Product Specialist in Integrated Solutions at Promega.
As sequencing technology races toward ever cheaper, faster and more accurate ways to read entire genomes, we find ourselves able to study biological systems at a level never before possible. From basic science to translational research, massively parallel sequencing (also known as next-generation sequencing or NGS) has opened up new avenues of inquiry in genomics, oncology and ecology.
Many commercial sequencing platforms have been established (e.g., Illumina, IonTorrent, 454, PacBio), and new technologies are developed every day to enable new and unique applications. However, all of these platforms and technologies work off the same general principle: nucleic acid must be extracted from a sample, arranged into platform-specific library constructs, and loaded into the sequencer. Regardless of the sample type or the platform used, every step throughout this workflow is critical for successful results. An often overlooked part of the NGS workflow is sample quantitation. Here we are presenting the first in a series of four short blogs about the critical step of quantitation in NGS workflows.
Sample input is critical to NGS in terms of both quality and quantity. Knowing how much DNA you have, often in nanogram quantities, can make the difference between success and failure. There are several key points in the NGS workflow where sample quantitation is important before you can proceed:
- After initial nucleic acid extraction from the sample matrix and before proceeding with library preparation
- Post-library preparation when pooling barcoded libraries for multiplexing
- Final pooled library quantitation immediately before loading for sequencing
There are several common methods for quantitating nucleic acids: UV-spectroscopy, Fluorescence spectoscopy, real-time quantitative PCR (qPCR). Because of inherent differences in sensitivity, specificity, time and cost, each of these techniques pose certain advantages and disadvantages with respect to the specific sample you are quantitating. Our next three blogs will discuss each of these methods against the backdrop of quantitating samples for NGS applications.