Scientists from the Wellcome Trust Sanger Institute recently published “An improved approach to mate-paired library preparation for Illumina sequencing,” a paper that came out in the open access journal Methods in Next Generation Sequencing.
The publication, from lead author Naomi Park, is the culmination of a project designed to establish an alternative to currently available commercial kits for mate-pair library construction. These existing methods of circularization “have differing limitations and are often linked to a single sequencing platform in a kit format, which may not be cost effective,” the authors write.
Mate-pair sequencing is often used for de novo genome assembly, detection of structural variants, genome finishing, and other projects in which long-range sequence data is beneficial. But preparing these libraries tends to be challenging — and gets even worse when there’s not much DNA to start with or when the DNA is degraded, the scientists note.
The paper compares a new method developed by the Sanger scientists with several existing mate-pair library construction approaches. “In order to generate unbiased and diverse Illumina mate-paired libraries containing even genomic sequence either side of a common adapter sequence, we altered the Illumina mate-pair protocol to use a modified SOLiD 4 hybridisation and ligation circularisation approach,” they write. Tests involving multiple libraries and sample types demonstrated that the new method increases library complexity and quality while reducing chimeras.
Most of the libraries involved in this study were size selected with BluePippin. “Alternative methods of size selection are amenable to this method, but may result in a differing breadth of size range and/or recovery,” the authors write. After comparing results from Pippin, manual gels, and bead size selection, they conclude, “The Blue Pippin provides the greatest recovery and accuracy of currently available commercial methods.”
Well, we’ve finally gone where no automated DNA size selection provider has gone before. Using compression-busting pulse-field programs and fancy gel formulations, we’ve tackled the large cut.
Our latest cassette type, BHF7510, comes with four cassette definitions:
1. High Range 40kb Marker W1- Tight
2. High Range 40kb Marker W1- Broad
3. High Range 50kb Marker W1- Tight
4. High Range 50kb Marker W1- Broad
In all four cases, the size selection is centered on the 40kb or 50kb target; range mode selections are not allowed. The “Tight” definitions provide very narrow distributions, while the “Broad” definition doubles the duration of elution to provide higher yields. Below is a summary of some our validation data.
Thanks to our crack R&D team for unlocking the mysteries of HMW fragments!
Figure 1. Data summary from High-Range cassette validation tests. Eluted DNA was run on agarose gels using the Pippin Pulse system (3-70kb preset program) and analyzed with the TL100 analysis software from Totallab, Ltd.
Field Inversion Gel Electrophoresis
We’ve set up this post to provide Pippin Pulse users with a resource to keep updated on new protocols and to post references that might be useful. We also invite users to feel use this page as a forum, if you see fit, and we’d enjoy showing interesting gel images and data (email these to email@example.com).
Our go-to book on theory:
“Electrophoresis of Large DNA Molecules:Theory and Applications” (E. Lai and B. Birren, eds), Current Communication in Cell & Molecular Biology Vol. 1,. Cold Spring Harbor Laboratory Press, New York, 1990
Here’s nice overview on pulsed field techniques:
Pulsed Field Electrophoresis for Separation of Large DNA
Download the Pippin Pulse user manual here
Our Gel Set-up
We use a Galileo Model 1214 RapidCastTM Mini Gel Unit running 12 X 14 cm gels using Lonza SeaKem® Gold Agarose. We use our Pippin Tris-TAPS buffers (which you can order directly from us in the US and Canada, Part No. KBB1001, under “Accessories” on our ordering page) or 0.5X TBE. The formulations can be found in the Pippin Pulse user manual or separately here.
Illumina released new sample prep protocol guidelines for generating mate pair libraries with its Nextera kit, and we’re pleased to report that the Pippin platform is the recommended choice for automated size selection.
You can check out the Nextera Mate Pair Sample Preparation Guide here . (We’re under Size Selection in Chapter 3, beginning on page 40 of the Guide.)
Illumina says that using an extra size selection step offers “more stringent” sizing than AMPure alone and lets users make libraries with larger fragments and more precise distribution than a gel-free approach. While the company has validated a manual approach in addition to the Pippin platform, Illumina’s guidelines note that “in our experience running a standard agarose gel does not provide as robust and reproducible results as the Sage Pippin Prep.”
In the user document, Illumina recommends the Pippin Prep with the 0.75% cassette and “eluting fragments with a broad range of sizes, of 3 to 6 kb in width, increasing in width with increasing fragment length (e.g. 2–5 kb, 4–8 kb or 6–12 kb).”
For current Pippin users, we would like to add that you can also use the 0.75% agarose dye-free cassette (BLF7510) with the BluePippin for equivalent results.
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!