Carl Linnaeus would be proud: A recent paper in PLoS One demonstrates the use of next-gen sequencing with genetic barcodes to accurately identify more than 100 different species from the Rodentia order. In the study, amplicons were run on the Pippin Prep from Sage Science to remove non-specific PCR products.
“Next-Generation Sequencing for Rodent Barcoding: Species Identification from Fresh, Degraded and Environmental Samples,” the paper from Maxime Galan, Marie Pagès, and Jean-François Cosson at the Center for Biology and Management of Populations at INRA, uses 454 GS-FLX sequencing. The authors note that correct species assignment in the diverse Rodentia order is quite challenging with morphological data alone.
In this work, the authors selected a 136 bp fragment from cytochrome b as a mini-barcode and then used it on more than 900 samples to determine its utility in accurately identifying species. Following a validation step, hundreds of samples of unknown identity were analyzed and the mini-barcode worked about 85 percent of the time, the scientists report. They also successfully tested degraded rodent DNA samples, including museum specimens and feces from rodent-eating predators.
The authors conclude, “This study demonstrates how this molecular identification method combined with high-throughput sequencing can open new realms of possibilities in achieving fast, accurate and inexpensive species identification.”
It’s that time of year again — we slow down at work, do some reflection, and gear up for a great (lucky?) ‘13….
In the field of genomics, 2012 was a fascinating year. On the NGS front, placements of desktop sequencers took off in terms popularity and performance, and the industry closed in on the $1000 genome. On the research front, studies on gene regulation continues to boggle the mind, and there seems to be an up-tick in interest in structural variation. This, and advances in cancer genomics, could very likely set off an avalanche of whole genome sequencing over the next few years.
For Sage Science, 2012 was a big year as well. We’ve established the Pippin as the go-to platform for high quality genomic libraries. And we launched BluePippin, the second in our Pippin automated size selection line, which features pulsed-field electrophoresis and collects fragments up to 50 Kb.
We were also delighted to see so many publications this year from scientists using the Pippin platform. Here are a few of our favorites:
• “Double Digest RADseq: An Inexpensive Method for De NovoSNP Discovery and Genotyping in Model and Non-Model
Species” from Peterson et al. in PLoS One
• “Towards quantitative metagenomics of wild viruses and other ultra-low concentration DNA samples: a rigorous assessment and optimization of the linker amplification method” from Duhaime et al. in Environmental Microbiology
• “Transcriptional Amplification in Tumor Cells with Elevated c-Myc” from Lin et al. in Cell
• “Synthetic Spike-in Standards Improve Run-Specific Systematic Error Analysis for DNA and RNA Sequencing” from Zook et al. in PLoS One
With that, we wish everyone a happy holiday season! See you in 2013.
Here at Sage Science, we are delighted to see more and more people signing up as customers of the Pippin platform. With so many instruments out in the wild, we thought it would be a good time to sit down with our customer service department (aka the incomparable Sadaf Hoda) to find out which topics are asked about most often, and what advice we can offer. Here’s what we came up with:
Q: My instrument came with a calibration fixture. What do I do with it?
A: It’s important to perform a simple LED calibration before each run of the Pippin instrument. This only takes 5 seconds and will give a pass/fail report letting you know that the LEDs are calibrated properly to optimize your run. When doing the calibration, be sure to center the fixture over the LED lights with the sticker facing up and the filter side facing down.
Q: The lid won’t close. Is something wrong with the instrument?
A: If you haven’t removed the tape strips from the buffer chambers on the cassette, that will prevent the electrodes from sitting down in the wells properly. Just take the tape strips off, and the lid should close fully.
Q: I stored the reagents and the cassettes separately, and now I can’t tell which reagents go with which cassettes. Help!
A: It’s very important to use the specific DNA marker or internal standard that is packaged with the cassette packages. We recommend that you store the cassettes at room temperature, and the reagents at 4oC. The labels on foil bags containing the cassette indicate which marker to use, as dp the cassette definitions in the software. For comprehensive information on cassettes go to our support page (www.sagescience.com/support) and download the Cassette Reference Chart for either the Pippin Prep or BluePippin. These are found in the “Guides” section.
Q: Are there different sample prep procedures for different cassettes?
A: The sample prep for cassettes is the same on the Pippin Prep and the BluePippin, but there are different protocols for cassettes with internal standards and ones with external markers. For internal standards, you will fill all five lanes with 40 microliters of sample (30 plus 10 microliters of standard/loading solution mix). For cassettes using external markers, you’ll fill one lane with 40 microliters of marker and the other four lanes with 40 microliters of sample (30 plus 10 microliters of loading solution).
Q: I’m using Illumina TruSeq kits. Does that have an effect on my size selection?
A: Yes, it’s been established that Illumina’s TruSeq kits require an offset for any size selection method, from manual gel excision to automated solutions like Pippin. Our experience with customers is that the offset is usually 100 to 150 base pairs if adaptor-ligated DNA is being run. The Illumina’s TruSeq user manual also provides guidelines about the offset to incorporate.
Q: I’ve finished my run. Now what?
A: We recommend that you immediately remove the cassette, and leave the lid open. If you leave the lid closed with the cassette still in the instrument, over time, salt can build up on the electrodes and lead to inaccurate sizing in one of the cassette lanes. This problem can be avoided by removing the cassette promptly and leaving the instrument lid open between runs.
Do you have a question that you feel should be answered here? Leave a reply, and we’ll post it!
Scientists at the University of Arizona, led by senior author and newly named Moore Foundation Investigator Matthew Sullivan, have published details of how to optimize sample prep methods for next-gen sequencing projects in which input DNA is negligible. As part of that work, the authors recommend the Pippin platform from Sage Science as the superior technology for automated size selection of libraries.
The paper, “Towards quantitative metagenomics of wild viruses and other ultra-low concentration DNA samples: a rigorous assessment and optimization of the linker amplification method,” from Duhaime et al., was published in September 2012 in Environmental Microbiology and is available via open access. The work focused on ways to reduce amplification bias, including precise DNA size selection, enzyme choice, and optimizing cycle number.
Sullivan and his team are using these methods for ocean-based viral metagenomics projects, but the methods translate to other projects that have very little DNA available. In this work, they looked at size selection in particular as a way to control size-related bias in the amplification step of sample preparation.
The paper reports on a comparison of Pippin to Solid Phase Reversible Immobilization (SPRI) from Beckman Coulter Genomics and to manual gels. The authors write:
“Of the three sizing fractionation methods tested for target recovery efficiency (fraction recovered DNA in target 400–600 bp size range), throughput (ease of applicability to numerous samples simultaneously), and risk of cross-sample contamination, Pippin Prep, an automated optical electrophoretic system that does not require gel extraction, was the most efficient and reproducible (94–96% of input DNA), with the tightest, most specific sizing.”
They note that SPRI was also high-throughput with low risk of contamination, but was the least efficient in recovery of the three methods tested, yielding 46-50 percent of the targeted size fraction after shearing.
“Based on this comparative analysis, we recommend the Pippin Prep automated electrophoretic system to prepare samples for [next-gen sequencing] libraries,” they conclude.
To learn more, read our case study here.
A collaboration among scientists at Monsanto and Bayer Crop Science has demonstrated the use of next-generation sequencing and a new bioinformatics method for analyzing the genome sequence of crops that have been genetically modified.
Lead authors David Kovalic and Carl Garnaat published their findings in The Plant Genome journal in a paper entitled “The Use of NexGen Sequencing and Junction Sequence Analysis Bioinformatics to Achieve Molecular Characterization of Crops Improved Through Modern Biotechnology.” You can view the full text here [PDF].
The goal of the project was to evaluate the use of next-gen sequencing compared to Southern blots and targeted sequencing of PCR products, the current standard for providing a molecular characterization of GM crops for both internal research and regulatory approval. The team studied strains of soybeans, both modified and wild type, using the Illumina platform to generate more than 75x coverage. During sample prep, they used the Pippin Prep from Sage Science to select a library with an average insert size of 280 base pairs.
As they write in the introduction:
“The overall strategy for this new molecular characterization method is to produce DNA sequence fragments that comprehensively cover the entire genome of test and control plants (i.e., the GM event under investigation, and the parent line from which it was derived) and use bioinformatic tools to analyze these DNA fragments. These bioinformatic analyses establish the insert/copy number and the presence/absence of backbone sequences.”
On the analysis front, the team developed a new bioinformatics approach called Junction Sequence Analysis to detect and characterize “novel chimeric sequences resulting from insertions into the native genome,” the authors report. In their assessment of the results, they add that “the method presented is capable of detecting complex events including those with multiple T-DNAs and sequence rearrangements.”
The scientists conclude that next-gen sequencing paired with Junction Sequence Analysis offers results equivalent to those from Southern blots — with the added advantages of “the simplicity, efficiency and consistency of the method,” they report, noting that their sequencing and analysis pipeline “provides a viable alternative for efficiently and robustly achieving molecular characterization of GM crops.”