Better (but not perfect) HMW DNA extraction protocol

I wrote some time ago about the protocol I used to prepare HMW DNA for the new HA412 assembly. The advantage of that protocol is that it doesn’t need much tissue to start with, it’s quick and can work quite well. However, it is also quite unreliable, and will sometimes fail miserably.

To prepare HMW DNA for H. anomalus I tried a different protocol, suggested by Allen Van  Deynze at UC Davis. They used it on pepper to prepare HMW DNA for 10X linked reads (the same application I had in mind), and obtained fragments of average size ~150-200 Kb. The resulting 10X assembly was quite spectacular (N50 = 3.69 Mbp for a 3.21 Gbp genome) and was recently published. Continue reading

Even-more-diluted BigDye

It turns out that you can use half the amount of BigDye that is recommended by NAPS/SBC for Sanger sequencing and have no noticeable drop in sequence quality. The updated recipe for the working dilutions:

BigDye 3.1 stock      0.5 parts

BigDye buffer            1.5 parts

Water                       1 part

I will prepare all future working dilutions using this recipe and put them in the usual box in the common -20ºC freezer. For more details on how to prepare a sequencing reaction see this post, and for how to purify them see this or this posts.

Streamlined GBS protocol

We already have a GBS protocol on the lab blog, but since it contains three different variants (Kate’s, Brook’s and mine) it can be a bit messy to follow. Possibly because I am the only surviving member of the original trio of authors of the protocol, the approach I used seems to have become the standard in the lab, and Ivana was kind enough to distill it into a standalone protocol and add some additional notes and tips. Here it is!

Simplified GBS protocol 2017

High Molecular Weight DNA

3rd generation sequencing technologies (PacBio, Oxford Nanopore) can produce reads that are several tens of Kb long. That is awesome, but it means that you also need to start from intact DNA molecules of at least that size. I thought the DNA extraction method I normally use for sunflower would be good enough, but that is not the case. Continue reading

BigDye 3.1

While cleaning up the freezer a few months back we found encrusted in a block of ice at the bottom of the common lab freezer a box with a seizable amount of BigDye 3.1, the reagent used to prepare samples for Sanger sequencing (basically a PCR mastermix with labelled nucleotides). As that is expensive stuff (what we have is worth 3-4000 dollars), I tested it to see if would still work. All the tests I did were rated “great sequence” when I got the results back from NAPS, so the BigDye is fine.

Thanks to a donation of buffer from NAPS, I have diluted some of the BigDye to the same working concentration NAPS uses (1 part of BigDye, 1.5 parts of buffer, 0.5 parts of water). Follow the instructions on the NAPS website ( to prepare your sample, and use 3 µl of the diluted BigDye. There are eight aliquots of about 150 µl each (50 reactions) in a box with a yellow “BigDye” label in the common -20 ºC freezer. If you plan to do only a few reactions at a time, consider making smaller aliquots for your personal use (BigDye doesn’t like repeated freeze-thaw cycles). To avoid confusion I kept the concentrated BigDye and dilution buffer in their Applied Biosystem box on the door of the first freezer on the right in the freezer room. If we run out of dilution just let me know, and I’ll be happy to prepare more.

Clean and cheap DNA from argophyllus (and other sunflowers)

Argophyllus has a reputation of being a plant it is hard to get DNA from. As a test for a larger project, I did a round of extractions from annuus, petiolaris and argophyllus. I used the modified 3% CTAB method I described before, starting from one fresh, frozen, very young (~1-2 cm long) leaf.

For all species, the CTAB extraction yielded about 50 µl of 200-500 ng/µl solution (10-25 µg in total) of clean (260/230 = 2.05-2.20) genomic DNA, with minimal shearing (see picture). Continue reading

DSN depletion for GBS libraries

This step is mentioned in our current GBS protocol, but I forgot to upload it until now. It’s basically the same as the WGS one, with minor changes. I am attaching a couple of bioanalyzer plots of the same library before and after DSN treatment. The sharp peaks/thick bands disappearing after the DSN treatment are likely chloroplast fragments.

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Home-brew WGS library multiplexing

There are two main ways to barcode WGS libraries so that they can be run together on a same lane:

– In-line barcodes: unique sequences are located at the very end of one or both adapters. This sequence will be at the very beginning of each read from a given library. This is the barcode system that is normally used fro GBS libraries as well.

– Indices: barcodes are in the middle of one or both adapters. These barcodes are read through an independent round of sequencing. For a paired-end library you would have therefore two rounds of sequencing of your fragment and a third round of sequencing for the index (and I guess a fourth one as well, if you have double indices). This is the system used in most commercial kits.

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Depletion of repetitive sequences – WGS libraries

As you know, the sunflower genome contains a large amount of repetitive sequences, that is why it is so big and so annoying to sequence. I have been working for a while on optimizing a depletion protocol, to try to get rid of some repetitive sequences in NGS libraries (transposons, chloroplast DNA…). Continue reading

Quick update: 96-well plate CTAB DNA extraction with fresh tissue

When I posted the protocol I have been using for CTAB DNA extraction in 96-well plates, I included results from a few plates I did starting from dried H. anomalus leaves I collected a couple of months earlier in Utah. While they showed that the method worked well enough when starting from “difficult” material, they were not exactly what you’d dream of when you decide to extract DNA, especially if you are starting instead from fresh material.

Here are the results from a plate of extraction I did starting from individual small (1.5-2 cm in length), young leaves from ~3 month-old H. anomalus plants. I collected the leaves directly in 96-well plates (I already put one metal bead in each well), put them on dry ice until I got to the lab (a couple of hours), left them overnight in the -80, and started extracting DNA the day after.


The final volume was 50 microliters, so total yield is for most samples between 10 and 30 micrograms of DNA. These are “real” DNA concentration measured by Qubit. Both average yield and purity are considerably higher than for dry tissue, and they are comparable to what you would get starting with frozen tissue using the single tube protocol (but you save a ton of time). Hope this gets you all more thrilled about 96-well plate DNA extractions 🙂

96-well plates CTAB DNA extraction

When I was working with Arabidopsis, 96-well CTAB DNA extraction was my best friend, and I spent many days extracting away tens of thousands of samples. Good times.

DNA extraction is much less pleasant in sunflower, but since I was reasonably happy with the results of single-tube 3% CTAB DNA extractions, I though I would try to scale it up to a 96-well plate format. Results of earlier attempts, with the participation of Brook and Cris, ranged from inconsistent to disastrous. Things all but improved when I tried again after coming back from Utah with a few hundreds dried samples. Since though the prospect of extracting them all one by one didn’t sound very attractive, I put some more effort into improving the protocol, and now it works quite nicely.

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Normalize/quantify your WGS libraries

Regardless of what method you use to make your Illumina libraries, if you added barcodes or indices you will need to normalize them before pooling (or otherwise have probably very uneven coverage). Or, you might want to know the exact molarity of your library before sending it for sequencing (although the need for that in our case is debatable, see later). The most accurate way to do both is probably by qPCR. Continue reading

(Probably the closest you can get to) Home-brew Illumina WGS libraries

As some of you might know, I have been working for the last few months on optimizing a protocol for Illumina WGS libraries that will reduce our dependency on expensive kits without sacrificing quality. The ultimate goal would be to be able to use WGS libraries as a more expensive but hopefully more informative alternative genotyping tool to GBS. Getting to that point ideally requires to develop:

1) A cheaper alternative for library preparation (this post)

2) A reliable multiplexing system (this other post)

3) A way to shrink the sunflower genome before sequencing it (because, as you know, it’s rather huge) (yet another post)

The following protocol is for non-multiplexed libraries. The protocol for multiplexed ones is actually identical, you just need to change adapters and PCR primers – more about that in the multiplexing post.

If you are planning to pool libraries and deplete them of repetitive elements, read carefully all three posts before starting your libraries (mostly because you might need to use different adapters and PCR primers)

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SPRI beads (almost) for free!

A few weeks ago Kristin posted a nice blog entry on home made SPRI beads, which effectively replace the ridiculously expensive commercial AMPure beads. (Editor’s note: see this post at RLR for more about the AMPure beads). Dan already ordered the ingredients to make them in our lab as well, and when they all arrived I volunteered to prepare and test them. Following are some results. If you don’t feel like reading through the whole post, the bottom line is that they work very well, and that you are very welcome to use them (they are in an aluminium-wrapped Falcon tube in the fridge, with a white “SPRI beads” label).

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