Category Archives: Science

What is ‘real-time’ phylogenomics?

Over the past few years I’ve been developing research, which I collectively refer to as ‘real-time phylogenomics’ – and this is the name of our mini-site for MinION-based rapid identification-by-sequencing. Since our paper on this will hopefully be published soon, it’s probably worth defining what I hope this term denotes now, what it does not – and ultimately where I hope this research is going.

‘Phylogenomics’ is simple enough, and Jonathan Eisen at UC Davis has been a fantastic advocate of the concept. Essentially, phylogenomics is scaled-up molecular systematics, with datasets (usually derived from a genome annotation and/or transcriptome) comprising many coding loci rather than a few genes. ‘Many’ in this case usually means hundreds, or thousands, so we’re typically looking at primarily nuclear genes, although organelles’ genomes may often be incorporated, since they’re usually far easier to reliably assemble and annotate. The aim is, basically to average phylogenetic signal over many loci by combining gene trees (or an analogous approach) to try and obtain phylogenies with higher confidence (single- or few-locus approaches, including barcodes no matter how judiciously chosen, capable of producing incorrect trees with high confidence). The process is intensive, since genomes must be sequenced and then assembled to a sufficient standard to be reasonably certain of identifying orthologous loci. This isn’t the only use of the term (which also refers to phylogenies produced from whole-genome metagenomics) but the most straightforward and common one as far as eukaryote genomics is concerned, and certainly the one uppermost in my mind.

However the results are often confusing, or at least more complex than we might hope: instead of a single phylogeny with high support from all loci, and robust to the model used, we often find a high proportion of gene trees (10-30%, perhaps) agree with each other, but not the modal (most common, e.g. majority rule consensus) tree topology. For instance among 2, 326 loci in our 2013 paper on phylogenomics of the major bat families, we found that position of a particular group of echolocators – which had been hotly debated for decades, based on morphological and single-locus approaches – showed such a pattern (sometimes supporting the traditional grouping of Microchiroptera + Megachiroptera, but over 60% of loci supporting the newer Yangochiroptera + Yinpterochiroptera system. This can be for a variety of reasons, some biological and some methodological. The point is that we have a sufficiently detailed picture to let us chose between competing phylogenetic hypothesis with both statistical confidence and intuition based on comparison.

These techniques have been on the horizon for a while (certainly since at least 2000) and gathered pace over the last decade with improvements in computing, informatics, and especially next-generation sequencing. The other half of this equation, ‘real-time’ sequencing, has emerged much more recently and centres, obviously, on the MinION sequencer. Most work using this so far has focused either on the very impressive potential long-read data offers for genomic analyses, particularly assembly, or rapid ID of samples e.g. the Quick/Loman Zika and Ebola monitoring studies; and our own work.

So what, exactly, do we hope to achieve with phylogenomic-type analyses using real-time MinION data, and why?

Well, firstly, our work so far has shown that the existing pipeline (sample -> transport -> sequence-> assemble genome-> annotate genes-> align loci-> build trees) has lots of room for speedups, and we’re fairly confident that the inevitable tradeoff with accuracy when you omit or simplify certain steps (laboratory-standard sequencing, assembly) is at least compensated for by the volume of data alone. Recall that a ‘normal’ phylogenomic tree similar to our bat one might take two or more postdocs/students a year to generate from biological samples, often longer. A process taking a week instead would let you generate something like 50 more analyses in a year! The most obvious application for this is just accelerating existing research, but the potential for transforming fieldwork and citizen science is considerable. This is because you can build trees that inform species relationships, even if the species in question isn’t known. In other words a phylogenome can both reliably identify an unknown sample, and also identify if it is a new species.

More excitingly, I think we need to have a deeper look at how we both construct and analyse evolutionary models. Life on earth can be accurately and fully described best by a network, not a bifurcating tree, but this applies to loci as well as single genes. In other words, there is a single network that connects every locus in every living thing. Phylogenetic trees are only a bifurcating projection of this, while single- or multi-locus networks only comprise a part.

We’ve hitherto ignored this fact, largely because (a) trees are often a good approximation, especially in the case of eukaryote nuclear genes, and (b) the data and computation requirements a ‘network-of-life’ analysis implies are formidable. However, cracks are beginning to appear, in both faces. Firstly, many loci are subject to real biological phenomena (horizontal gene transfer, selection leading to adaptive convergence, etc) which give erroneous trees as discussed above. Meanwhile prokaryotic and viral inference is rarely even this straightforward. Secondly, expanding computing power, algorithmic complexity, and sequencing capacity (imagine just 1,000 high schools across the world, regularly using a MinION for class projects…) mean the question for us today really isn’t ‘how do we get data’, but ‘how ambitious do we want to be with it?’

Since my PhD but especially since 2012, I’ve been developing this theme. Ultimately I think the answer lies in the continuous analysis of public phylogenomic data. Imagine multiple distributed but linked analyses, continuously running to infer parts of the network of life, updating their model asynchronously both as new data flood in, and by exchanging information with each other. This is really what we mean by real-time phylogenomics – nothing less than a complete Network of Life, living in the cloud, publicly available and collaboratively and continuously inferred from real-time sequence data.

So… that’s what I plan to spend the 2020s doing, anyway.

 

Some aspects of BLASTing long-read data

Quick note to explain some of the differences we’ve observed working with long-read data (MinION, PacBio) for sample ID via BLAST. I’ll publish a proper paper on this, but for now:

  • Long reads aren’t just a bit longer than Illumina data, but two, three, four or possibly even five orders of magnitude longer (up to 10^6 already, vs 10^2). This is mathematically obvious, but extremely important…
  • … the massive length means lots of the yield is in comparatively few reads. This makes yield stats based on numbers of reads positively useless for comparison with NGS. Also…
  • Any given long read contains significantly more information than a short one does. Most obviously the genomics facilities of the world have focused on their potential for improving genome assembly contiguity and repeat spanning (as well as using synteny to spot rearrangements etc) but we’ve also shown (Parker et al, submitted) that whole coding loci can be directly recovered from single reads and used in phylogenomics without assembly and annotation. This makes sense (a ~kb long read can easily span a whole gene, also ~kb in scale) but it certainly wasn’t initially obvious, and given error rates, etc, it’s surprising it actually works.
  • Sample ID using BLAST actually works very differently. In particular, the normal ‘rubbish in, rubbish out’ rule is inverted. In other words, nanopore reads (for the time being) may be long, but inevitably contain errors. However, this length means that assuming BLAST database sequences are approximately as long/contiguous, Nanopore queries tend to either match database targets correctly, with very long alignments (hundreds/thousands of identities), or not at all.

This last point is the most important. What it means is that, for a read, interpreting the match is simple – you’ll either have a very long alignment to a target, or you won’t. Even when a read has regions of identity to more than one species, the correct read has a much longer cumulative alignment length overall for the correct one. This is the main result of our paper.

The second implication is that, as it has been put to me, for nanopore reads to be any good for an ID, you have to have a genomic database. While this is true in the narrow sense, our current work (and again, this is partly in our paper, and partly in preparation) shows that in fact all that matters is for the length distribution in the reference database to be similar to the query nanopore one. In particular, we’ve demonstrated that a rapid nanopore sequencing run, with no assembly, can itself serve as a perfectly good reference for future sample ID. This has important implications for sample ID but as I said, more on that later 😉

Science and (small) business

Over the last 10-20 years there’s been a revolution in academic science (or should that be ‘coup’?) where many aspects of the job have been professionalised and formalised, especially project management but management in general. This generally includes tools like GANTTs, milestones, workload models, targets and many other things previously unmentionable in academia but common in industry, especially large organisations. Lots of academics will tell you they think it’s bureaucratic overkill, intrusive, a waste of time, and worse (to put it mildly) but the awkward truth is that, as lab groups steadily increased in size (as fewer, larger grants went to increasingly senior PIs or consortia) many of the limitations of the collegiate style of the past, centred on a single academic with a tight-knit group, have been exposed.

Frequently the introduction of ‘management practices’, often after hiring expensive consultants, is accompanied by compulsory management training. Sometimes it can be an improvement. More normally (in my experience) whether an improvement in outcomes (as distinct from ‘efficiency’) has been achieved probably depends on whether you cost in staff time (or overtime) and morale. You can make arguments either way.

But I can’t help thinking: why are we attempting to replicate practices from big/massive private sector organisations, anyway? I suspect, the answer in part is because those are the clients management consultants have the most experience working with. More seriously, those organisations differ in fundamental respects from even the largest universities, let alone individual research projects. This is because large companies:

  • Add value to inputs to create physical goods or services that are easily costed by the market mechanism (this is the big one)
  • Usually have large cash reserves, or easy access to finance (tellingly when this ends they usually get liquidated)
  • Keep an eye on longer-term outcomes, but primarily focus on the 5-10 year horizon
  • Compete directly with others for customers (in some respects an infinite resource)
  • Are answerable, at least, yearly, to shareholders – with share value and dividends being the primary drivers of shareholder satisfaction.

Meanwhile, universities (and to an even greater extreme, research groups/PIs):

  • Produce knowledge outputs with zero market value*
  • Live hand-to-mouth on short grants
  • Need long-term, strategic thinking to succeed (really, this is why we get paid at all)
  • Compete indirectly for finite resources grants and publications, based partly on track record and partly on potential.
  • Answer, ultimately, to posterity, their families, and their Head Of Department

I want to be clear here – I’m not saying, by any means, that previous management techniques (ie, ‘none’) work well in today’s science environment – but I do think we should probably look to other models than, say General Motors, or GlaxoSmithKline. The problem is often compounded because PIs have no business experience (certainly not in startups) while consultants often come from big business – their ability to meet in the middle is limited.

Instead small and medium enterprises (SME)s are a much closer model to how science works. Here good management of resources and people is extremely important, but the scale is much smaller, permitting different management methods, often focussing on flexibility and results, not hierarchies and systems. For instance, project goals are often still designed to be SMART (specific, measurable, achievable, realistic, time-scaled) but these will be revisited often and informally, and adjusted whenever necessary. Failure is a recognised part of the ongoing process. This is the exact opposite to how a GANTT, say, is used in academia: often drawn up at the project proposal (design) stage, it is then ignored until the end of the grant, when the PI scrabbles to fudge the outcomes, goals, or both to make the work actually carried out fit, so they don’t get a black mark from the funder/HoD for missing targets.

There are plenty of other models, and they vary not just by organisation size/type (e.g. tech startup, games studio, SkunkWorks, logistics distributor, niche construction subcontractor) but you see what I mean: copying ‘big business’ wholesale without looking at the whole ecosystem of business practices makes little sense…

*Obviously not all, or even most, scientific output will never realise any economic value – but it can be years, or centuries removed from the work to create it. And spin-outs are relevant to a tiny proportion of PI’s work, even in applied fields.

Single-molecule real-time (SMRT) Nanopore sequencing for Plant Pathology applications

A short presentation to the British Society for Plant Pathology’s ‘Grand Challenges in Plant Pathology’ workshop on the uses of real-time DNA/RNA sequencing technology for plant health applications.

Doctoral Training Centre, University of Oxford, 14th September 2016.

Slides [SlideShare]: cc-by-nc-nd


More MinION – the ‘1D rapid’ prep

My last MinION post described our first experiments with this really cool new technology. I mentioned then that their standard library prep was fairly involved, and we heard that the manufacturers, Oxford Nanopore, were working on a faster, simpler library prep. We got in touch and managed to get an early prototype of this kit for developers*, so we thought we’d try it out. So our** experiments had three aims:

  • Try out this new rapid kit
  • Try out different extraction methods, to see how they worked with the kit
  • See if we could sequence some fairly damaged DNA with the kit

This is a lot of combinations to perform over a few days on one sequencing platform! Our experience from the last run gave us hope we could manage it all (we did, but with a lot of headaches over disk space – it turns out one drawback of being able to run multiple concurrent sequencing runs is hard-drive meltdown, unless you’re organised from the start – oops). In fact, to keep track of all the reads we added an ‘index’ function to the Poretools package. I really recommend you use this if you’re planning your own work.

We had eight samples to sequence, a 15-year-old dried fungarium specimen of Armillaria ostoye (likely to be poor quality DNA; extracted by Bryn Dentinger with a custom technique); some fresh Silene latifolia (Qiagen-extracted, which we’d used successfully with the previous, ‘2D’ library prep); and six arbitarily-selected plant samples, both monocots and dicots, extracted by boiling with Chelex beads (more about them at the end).

IMG_4555

First we prepared normal ‘2D’ libraries from the Silene and Armillaria. These performed as expected from our December experiments, even the Armillaria giving decent numbers and lengths of reads (though not as many as we hoped, with some indication of worse Q-scores. We put this down to nicks in the fungarium DNA, and moved on to the 1D preps while the sequencer was still running.

The ‘1D’ in the rapid kit (vs ‘2D’ in the normal kit) refers to how many DNA strands are sequenced; in the 2D version, both forward and reverse-complement strands are sequenced. This is slower to prepare (extra adapters etc to link the two strands for sequencing) and also runs through the MinION more slowly (twice as much DNA, plus a hairpin moment) but is roughly twice as accurate, since each base is read twice. The 1D kit, on the other hand, results in single-stranded fragments, meaning we could expect lower accuracy traded off for higher speed. And the rapid kit really was fast – starting from purified extracted DNA, we added all the necessary adapters for sequencing in well under 15mins, ready to sequence.

lenQuals
The sequencing itself (for both the Armillaria and the Silene) went spectacularly well. Remember, this is unsheared genomic DNA, and imagine our surprise when we started to see 25, then 50, then 100, then 150kb reads come off the sequencer – many mapping straight away to reference genomes! It turns out that the size distribution of the 1D prep is much more long-tailed than the 2D/g-TUBE one. In fact, whereas the the 2D library looks like a normal-ish gamma, the 1D reads are more like an inverse exponential – lots of short stuff and then a very long tail with some mega-whopper reads in. Reads so long, in fact, that mapping them the same way as Illumina short-read data would be a bit bonkers…

As for accuracy, well, the Q-scores are definitely lower in the 1D prep; around half the 2D as we expected. On the other hand, they were still matching reference databases via BLAST/BLAT/BWA quite happily – so if your application was ID, who cares? Equally, combining mega-long 1D reads with more shorter but accurate reads could be a good way to close gaps in a de novo genome sequencing project. One technical point – there is definitely a lot more relative variation in the Q-scores for short (<1000bp) reads than for longer ones: the plot above shows the absolute difference in mean Q-scores in (first – vs – second) halves of a subset of 2,300 1D reads. You can see that below 1kb, Q-score variation exceeds 4 (bad news when mean is about there) while longer reads have no such effect (quick T-test confirms this).

So in short, the 1D prep is great if you just want to get some DNA on your screen ASAP, and/or long reads to boot. In fact, if you came up with a way to size-select all the short gubbins out before sequencing, you’d have one mega-cool sequencing protocol! What about the last bit of our test – seeing if a quick and dirty extraction could work, too? The results were… mixed-to-poor. Gel electrophoresis and Qubit both suggested the extracted DNA was pretty poor quality/concentration, and if we didn’t believe them, the gloopy, aromatic, multicoloured liquid in the tubes supplied direct evidence to our eyes. So rather than test those samples first (and risk damaging a perfectly good flowcell early on in the experiment), we held them back until the end when only a handful of pores were left. In this condition it’s hard to say whether they worked, or not: the 50 or so reads we got over an hour or two from fewer than 10 pores is a decent haul, and some of them had some matches to congenerics in BLAST, but we didn’t really give them a full enough test to be sure.

Either way, we’ll be playing around with this more in the months to come, so watch. This. Space…

 

*Edit: Oxford Nanopore have recently announced that the rapid kit will be out in April for general purchase.

**Again working, for better or worse, with fellow bio-beardyman Dr. Alex Papadopulos. Hi Alex! This work funded by a Royal Botanic Gardens, Kew Pilot Study Grant.

Copying LOADS of files from a folder of LOADS *AND* LOADS more in OSX

Quick one this, as it’s a tricky problem I keep having to Google/SO. So I’m posting here for others but mainly myself too!

Here’s the situation: you have a folder (with, ooh, let’s say 140,000 separate MinION reads, for instance…) which contains a subset of files you want to move or copy somewhere else. Normally, you’d do something simple like a wildcarded ‘cp’ command, e.g.:

1
host:joeparker$ cp dir/*files_I_want* some/other_dir

Unfortunately, if the list of files matched by that wildcard is sufficiently long (more than a few thousand), you’ll get an error like this:

1
-bash: /bin/cp: Argument list too long

In other words, you’re going to have to be more clever. Using the GUI/Finder usually isn’t an option either at this point, as the directory size will likely defeat Finder, too. The solution is pretty simple but takes a bit of tweaking to work in OSX (full credit to posts here and here that got me started).

Basically, we’re going to use the ‘find’ command to locate the files we want, then pass each one in turn as an argument to ‘cp’ using ‘find -exec’. This is a bit slower overall than doing the equivalent as our original wildcarded command but since that won’t work we’ll have to lump it! The command* is:

1
find dir -name *files_I_want* -maxdepth 1 -exec cp {} some/other_dir \;</p>

Simple, eh? Enjoy :)

*NB, In this command:

  • dir is the filesystem path to start the search; ‘find’ will recursively traverse the directory tree including and below this folder;
  • -name *glob* gives the files to match;
  • -maxdepth is how deep to recurse (e.g. 1 = ‘don’t recurse’);
  • cp is the command we’re executing on each found file (could be mv etc);
  • {} is the pipe argument standing for the found file;
  • some/other_dir is the destination argument to the command invoked by -exec

How to fake an OSX theme appearance in Linux Ubuntu MATE

I’ve recently been fiddling about and trying to fake an OSX-style GUI appearance in Linux Ubuntu MATE (15.04). This is partly because I prefer the OSX GUI (let’s be honest) and partly because most of my colleagues are also Mac users mainly (bioinformaticians…) and students in particular fear change! The Mac-style appearance seems to calm people down. A bit.

The specific OS I’m going for is 10.9 Mavericks, because it’s my current favourite and nice and clear. There are two main things to set up: the OS itself and the appearance. Let’s take them in turn.

1. The OS

I’ve picked Ubuntu (why on Earth wouldn’t you?!) and specifically the MATE distribution. This has a lot of nice features that make it fairly Mac-y, and in particular the windowing and package management seem smoother to me than the vanilla Ubuntu Unity system. Get it here: https://ubuntu-mate.org/vivid/.* The installation is pretty painless on Mac, PC or an existing Linux system. If in doubt you can use a USB as the boot volume without affecting existing files; with a large enough partition (the core OS is about 1GB) you can save settings – including the customisation we’re about to apply!

*We’re installing the 15.04 version, not the newest release, as 15.04 is an LTS (long-term stable) Ubuntu distribution. This means it’s supported officially for a good few years yet. [Edit: Arkash (see below) kindly pointed out that 14.04 is the most recent LTS, not 15.04. My only real reason for using 15.04 therefore is ‘I quite like it and most of the bugs have gone'(!)]

2. The appearance

The MATE windowing system is very slick, but the green-ness is a bit, well, icky. We’re going to download a few appearance mods (themes, in Ubuntu parlance) which will improve things a bit. You’ll need to download these to your boot/install USB:

Boot the OS

Now that we’ve got everything we need, let’s boot up the OS. Insert the USB stick into your Mac-envious victim of choice, power it up and enter the BIOS menu (F12 in most cases) before the existing OS loads. Select the USB drive as the boot volume and continue.

Once the Ubuntu MATE session loads, you’ll have the option of trialling the OS from the live USB, or permanently installing it to a hard drive. For this computer I won’t be installing to a hard drive (long story) but using the USB, so customising that. Pick either option, but beware that customisations to the live USB OS will be lost should you later choose to install to a hard drive.

When you’re logged in, it’s time to smarten this baby up! First we’ll play with the dock a bit. From the top menu bar, select “System > Preferences > MATE Tweak” to open the windowing management tool. In the ‘Interface’ menu, change Panel Layouts to ‘Eleven’ and Icon Size to ‘Small’. In the ‘Windows’ menu, we’ll change Buttons Layout to ‘Contemporary (Left)’. Close the MATE Tweak window to save. This is already looking more Mac-y, with a dock area at the bottom of the screen, although the colours and icons are off.

Now we’ll apply some theme magic to fix that. Select “System > Preferences > Look and Feel > Appearance”. Now we can customise the appearance. Firstly, we’ll load both the ‘Ultra-Flat Yosemite Light’ and ‘OSX-MATE’ themes, so they’re available to our hybrid theme. Click the ‘Install..’ icon at the bottom of the theme selector, you’ll be able to select and install the Ultra-Flat Yosemite Light theme we downloaded above. It should unpack from the .zip archive and appear in the themes panel. Installing the OXS-MATE theme is slightly trickier:

  • Unzip (as sudo) the OSX-MATE theme to /usr/share/themes
  • Rename it from OSX-MATE-master to OSX-MATE if you downloaded it from git as a whole repository (again, you’ll need to sudo)
  • Restart the appearances panel and it should now appear in the themes panel.

We’ll create a new custom theme with the best bits from both themes, so click ‘Custom’ theme, then ‘Customise..’ to make a new one. Before you go any further, save it under a new name! Now we’ll apply changes to this theme. There are five customisations we can apply: Controls, Colours, Window Border, Icons and Pointer:

  • ControlsUltra-Flat Yosemite Light
  • Colours: There are eight colours to set here. Click each colour box then in the ‘Colour name’ field, enter:
    • Windows (foreground): #F0EAE7 / (background): #0F0F0E
    • Input boxes (fg): #FFFFFF / (bg): #0F0F0E
    • Selected items (fg): #003BFF / (bg): #F9F9F9
    • Tooltips: (fg): #2D2D2D / (bg): #DEDEDE
  • Window borderOSX-MATE
  • IconsFog
  • PointerDMZ (Black)

Save the theme again, and we’re done! Exit Appearance Preferences.

Finally we’ll install Solarized as the default terminal (command-line interface) theme, because I like it. In the MATE Terminal, Unzip the solarized-mate-terminal archive, as sudo. Enter the directory and simply run (as sudo) the install script using bash:


$ sudo unzip solarized-mate-terminal
$ cd solarized-mate-terminal
$ bash solarized-mate.sh

Close and restart the terminal. Hey presto! You should now be able to see the light/dark Solarized themes available, under ‘Edit > Profiles’. You’ll want to set one as the default when opening a new terminal.

Finally…

Later, I also installed Topmenu, a launchpad applet that gives an OSX-style top-anchored application menu to some linux programs. It’s a bit cranky and fiddly though, so you might want to give it a miss. But if you have time on your hands and really need that Cupertino flash, be my guest. I hope you’ve had a relatively easy install for the rest of this post, and if you’ve got any improvements, please let me know!

Happy Tweaking…

Messing about with the MinION

IMG_4543Molecular phylogenetics – uncovering the history of evolution using signals in organisms’ genetic sequences – is a powerful science, the latest expression of the human desire to understand our common origins. But for all its achievements, I’d always felt something was, well lacking from my science. This week we’ve been experimenting with the MinION for the first time, and now I know what that is: immediacy. I’ll return to this theme later to explain how exciting this realisation is, but first we better ask:

What is a MinION?

NGS nerds know about this already, of course, but for the rest of us: The MinION (minh-aye-on) is a USB-connected device marketed by a UK company, Oxford Nanopore, as ‘a portable real-time biological analyser’. Yes – that does sound a lot like a tricorder, and for good reason: just like the fictional device, it promises to make the instant identification of biological samples a reality. It does this using a radically different new way to ‘read’ DNA sequences from a liquid into a letters (‘A, C, G, T‘) on a computer screen. Essentially, individual strands of DNA are pulled through a hole (a ‘nanopore’) in an artificial membrane, like the membranes that surround every living cell. When an electric field is applied to the membrane, the individual DNA letters (actually, ‘hexamers’ – 6-letter chunks) passing through the nanopore can be directly detected as fluctuations in the field, much like a magnetic C-60 cassette tape is read by a magnetic tape head. The really important thing is that, whereas other existing DNA-reading (‘sequencing’) machines take days-to-weeks to read a sample, the MinION produces output in minutes or even seconds. It’s also (as you can see in the picture, above) a small, wait tiny device.

Real-time

This combination of fast results and small size makes the tricorder dream possible, but this is more than just a gadget. Really, having access to biological sequences at our fingertips will completely change biology and society in many ways, some of which Yaniv Erlich explored in a recent paper. In particular, molecular evolutionary biologists will soon be able to interact with their subject matter in ways that other scientists have long taken for granted. What I mean by this is that in many other empirical disciplines, researchers (and schoolkids!) are able to directly observe, or easily measure, the phenomena they study. Paleontologists dig bones. Seismologists feel earthquakes. Zoologists track lions. And so on. But until now, the world of genomic data hasn’t been directly observable. In fact reading DNA sequences is a slow, expensive pain in the arse. And in such a heavily empirical subject, I can’t help but feel this is a hindrance – we are burdened with a galaxy of baroque models to explain variations in the small number of observations we’ve made, and I’ve got a hunch more data would actually consign many of them to the dustbin.

Imagine formulating, testing, and validating or discarding genetic hypotheses as seamlessly as an ecologist might survey a new forest…

IMG_4531

Actual use

That’s the spiel, anyway. This week we actually used the MinION for the first time to sequence DNA (I’d been running some other technical tests for a couple of months, but this was our first attempt with real samples) and since it seems lots more colleagues want to ask about this device than have had a chance to use it, I thought I’d share our experiences. I say ‘we’ – work took place thanks to a Pilot Study Fund grant at the Royal Botanic Gardens, Kew, in collaboration with Dr. Alex Papadopulos (and really useful input from Drs. Andrew Helmstetter, Pepijn Kooij and Bryn Dentinger). It’s worth pointing out that although there’s a lot of hype about the device, it is still technically in a prototype/public-beta type phase, so things are changing all the time in terms of performance, etc.

IMG_4547First up, the size. The pictures don’t really do it justice… compared with existing machines (fridge-sized, often) the MinION isn’t small, it’s tiny. The thing itself, plus box and cable, would easily fit into a side pocket on any laptop bag. So the ‘portable’ bit is certainly true, as far as the platform itself is concerned. But there’s a catch – to prepare a biological sample for sequencing on the MinION, you first have to go through a fairly complicated set of lab steps. Known collectively as ‘library preparation’ (a library here meaning a test tube containing a set of DNA molecules that have been specially treated to prepare them to be sequenced), the existing lab protocol took us several hours the first time. Partly that was due to the sheer number of curious onlookers, but partly because some of the requisite steps (bead cleanups etc) just need time and concentration to get right. None of the steps is particularly complicated (I’m crap at labwork and just about followed along) but there’s quite a few, and you have to follow the steps in the protocol carefully.

Performance

So how did the MinION do? We prepared two libraries; one from a control sample of bacteriophage lambda (a viral genome, used to check the lab steps are working) and another from Silene latifolia, a small flowering plant and one of Alex’s faves. The results were exciting. In fact they were nerve-wracking – initially we mixed up the two samples by mistake (incredible – what pros…) and were really worried when, after a few hours’ sequencing, not a single DNA read matching the lambda genome had appeared. After a lot of worrying, and restarting various analyses, including the MinION itself, we eventually realised our mistake, reloaded the MinION with the other sample and – hey presto! – lambda reads started to pour out of the software. In the end, we were able to get a 500x coverage really quickly (see reads mapped to the reference genome, below):

lambda

You can see from the top plot that we got good even coverage across the genome, while the bottom plot shows an even (ish) read length distribution, peaking around 4kb. We’d tried for a target size of 6kb (shearing using g-TUBEs), so it seems the actual output read size distribution is lower than the shearing target – you’d need to aim higher to compensate. Still, we got plenty of reads longer than 20, or even 30,000 base-pairs (bp) – from a 47kbp genome this is great, and much much much longer than typical Illumina paired-end reads of ~hundreds of bp.

Later on the next day, we decided we’d done enough to be sure the library preparation and sequencing were working well, so we switched from the lambda control to our experimental (S.latifolia) sample. Again, we got a good steady stream of reads, and some were really long, over 65,000bp. Crucially, we were able to BLAST these against NCBI and get hits against the NCBI public sequence database and get Silene hits straight back. We were also able to map them onto the Silene genome directly using BWA, even with no trimming or masking low-quality regions. Overall, we got nearly a full week’s sequencing from our flowcell, with ~24,000 reads at a mean length of ~4kb. Not bad, and we’d have got more if we hadn’t wasted the best part of the run sodding about while we troubleshot our library mix-up at the start (the number of active pores, and hence sequencing throughput, declines over time).

The bad: There is a price for these long, rapid reads. Firstly, the accuracy is definitely lower than Illumina – although the average lambda library sequencing accuracy in our first library was nearly 90%, on some of our mapped Silene reads it dropped much lower – 70% or so. This has been well documented, of course. Secondly, the *big* difference between the MinION in use and a MiSeq or HiSeq is the sheer level of involvement needed to get the best from the device – whereas both Illumina machines are essentially push-button operated, the MinION seems to respond well if you treat it kindly (reloading etc) and not if you don’t (introducing air bubbles while loading seemed to wreck some pores)

The very, very, good: Ultimately though, we came away convinced that these issues won’t matter at all in the long run. ONT have said that the library prep and accuracy are both boing improved, as are flowcell quality (prototype, remember). Most importantly the MinION isn’t really a direct competitor to the HiSeq. It’s a completely different instrument. Yes, they both read DNA sequences, but that’s where the similarity ends. The MinION is just so flexible, there’s almost an infinite variety of uses.

In particular, we were really struck that the length of the reads means simple algorithms like BLAST can get a really good match from any sample with just a few hundreds, or tens, of reads from a sample. You just don’t need millions of 150bp reads to match an unknown sample to a database with reads this long! Coupled with the fact that the software is real-time and flowcells can be stopped and started, and you have the bones of a really capable genetic identification system for all sorts of uses; disease outbreaks (in fact see this great work on Ebola using MinION); customs control of endangered species; agriculture; brewing – virtually anything, in fact, where finding out about living organisms’ identity or function is needed.

The future.

There’s absolutely no question at all: really, these devices are the future of biology. Maybe the MinION will take off (right now, the buzz couldn’t be hotter), or a competitor will find a way to do even more amazing things. It doesn’t matter how it comes about, though – the next generation of biologists really will be living in a world where, in 10, 15, 20 years, at most, the sheer ubiquity of sequencers like this will mean that most, if not all eukaryotes’ genomes will have been sequenced, and so can be easily matched against an unknown sample. If that sounds fantastic, consider that the cumulative sequencing output of the entire world in 1988 amounted to a little over 100,000 sequences, a yield equivalent to a single good MinION run. And while most people in 1988 thought that, since the human genome might take 30 or 40 years to sequence, there was little point in even starting, a few others looked around at the new technology, its potential, and drew a different conclusion. The rest is history…