My name is Peter Rendbæk, and I’m currently a master student in the Albertsen lab. The overarching aim of my master project, is as a pre-test for several of the new big projects in the group, which focus on applying the on-line bacterial identification for process control at wastewater treatment plants. Hence, last couple of months I have been working on the project “Developing methods for on-site DNA sequencing using the Oxford Nanopore MinION platform”. The MinION has improved a lot since its release three years ago, and it can now be used to make rapid determination of bacterial compositions.
The potential for this fast and mobile DNA-sequencing is mind-blowing. However, given that the technology is here now (!), there has been relatively little focus on portable, fast, easy and robust DNA extraction. Hence, I’ve spent the last months on trying to develop a fast, cheap, mobile, robust and easy to use DNA extraction method.
There is a significant amount of bias connected with DNA extraction, but the bias associated with wastewater treatment samples has been investigated in depth. However, the “optimized method” is not suited for on-site DNA-extraction. There are 3 principle steps in DNA extraction, cell lysis, debris removal and DNA isolation, which I will cover below and discuss how I simplified each step.
In general, complex samples require bead beating for cell lysis and homogenization. The problem is that our in-house bead-beating is done by a big table top tool weighing 17 kg, which makes it hard to transport. However, I came across a blog post from loman labs about sample preparation and DNA extraction in the field for Nanopore sequencing. In the blog post, the possibilities of a portable bead beater outlined, by the use of a remodeled power-tool. I thought this was interesting, so I went out and bought an Oscillating Multi-Tool cutter and tried this with lots of duct tape…
The amazing part was that it worked! But the problem was that the samples would get “beaten” differently depend on how you taped the sample to the power-tool, which could give rise to variation large variations in the observed microbial community.
I solved this by 3D printing an adapter to the power-tool that fits the bead-beater tube (Finally, a good excuse to use a 3D printer!). I used Solidworks to design the adapter and collaborated with our local department of mechanical and manufacturing engineering (m-tech) in 3D printing it. You can make your own by simply downloading my design from Thingiverse (It did take a few iterations to make it durable enough, and I still use a little duct tape..).
After the bead beating, the cell debris removal is done by centrifugation. Our “standard” protocol recommends centrifugation at 14000 x G for 10 minutes at 4 C. However, in our minds that seemed a little extensive and requires a huge non-transportable centrifuge… Alternatively, there are a lot of possibilities to use small, easy to transport and easy to use centrifuges if we do not have to centrifuge at 14.000 xG at 4 C. There is even the possibility to 3D print a hand-powered centrifuge. However, I did not follow this path, as it seems a bit dangerous… After several tests, we discovered that a simple table top centrifuge could do the job perfectly well, using 2000 xG for 1 min at room temperature if we combined it with the DNA isolation described below.
The last step is DNA isolation, I tried several different methods, but we got the idea to simply use Agencourt AMPure XP that is routinely used in e.g. PCR purification (we 10 diluted the AMPure XP beads 1:10 to save some money and it seems to work just as good). And… It works..
So, now you have an overview of the method I developed. The most amazing part is that it works! It takes 10-15 minutes from the sample is taken until you’ve got ready DNA for use, compared to 60+ minutes for our “standard” protocol. Furthermore, it requires inexpensive equipment that can be carried in a small suitcase. So, just to prove that this approach is fast, I filmed myself doing the DNA extraction with a GO-PRO camera, as you can see below.
The next part is to test the MinION in the lab. How, fast can we identify bacteria and is the extracted DNA compatible with the downstream library preparation, which we hope to do on the our new and shiny Voltrax (which is now moving liquids!).
This is an absolutely awesome post ! Thank you for sharing !
I was curious about what you call “pure” DNA: what are the typical 260/280 and 260/230 ratios that you get through this methodology ?
Hi, Anne-Lise thank you for the question.
I measured the sample that got extracted in the video using a Nanodrop1000 (Thermo Fisher Scientific). The 260/280 ratio DNA is 1.80, but the 260/230 ratio was 1.65 (see picture below). Ideally it should be at 2.0-2.2 and the low ratio indicate some level of organic contamination. It should be noted, that we have not tried to use the extracted DNA on the MinION yet, and one of the criteria for the Oxford Nanopore “Rapid kit” is a purity at OD 260/230 at 2.0-2.2… However, we were able to perform standard 16 rRNA PCR amplification and Illumina sequencing on the extracted DNA.
Thanks Peter !
I am amazed by your ratios: they’re really good for such a quick procedure !
Indeed RAD002 is really sensitive to the purity of samples. I tried it with some gDNA from a fungi isolate. 260/230 ratio was <2 and it has been a complete bust. I will not use it anymore useless I am 100% sure of being in 2.0-2.2.
Hi Peter, I read your article but i missed something about the size of your ‘isolated’ DNA fragments. The bigger the better? for sequencing with the minion? Did you look at it?
Since Peter left our lab after finishing his masters degree I will provide you with the answer. The peak was at 16-17 kbp. Peter determined this using a genomic screentape. The size distribution can be seen below:
its good to see the multitool bead beater. I had a similar idea for 18V tools , mutltitooll for lysis and angle grinder modified to be a centrifuge. Also been thinking about modifying a fidget spinner as a simple centrifuge to get everything to the bottom of the tube, would be interesting to see if it can elute a sample from a spin column.
all the best