Normalisation made simple...please?

We do lots of DNA, RNA and NGS library normalisations mostly by hand after calculating dilutions in Excel. It works well but is cumbersome and prone to error. We're fortunate to have an Agilent Bravo robot and are now automating more of this with excellent results. The most obvious impact has been on NGS library pooling, below is our most recent run where the variability on all but 9 libraries is pretty much perfect.

However automation is a pain and it would be great if there were a solution for normal labs. We've discussed an idea with pipette companies in the past but no-one appears to offer anything yet so I thought I'd suggest it here as well. If you've seen something on the market please do comment and let me know (Sartorius are almost there with the eLine pipette see link below).

If you like the idea follow up with you favourite pipette rep and lets see if someone can make one!

The background to the idea: normalisation can be done by combining a standard volume of sample with a know volume of water or diluent. Any lab trying to do this manually is almost certainly taking concentration measurements from PicoGreen or qPCR and working out how much water to add to each well of a 96well plate with sample already added.

The idea: We use repeat pipettes for may applications and there do exist programmable versions. Repeat pipettes are great for adding the same volume of a master mix to all wells of a plate and several companies make them. We use the slightly older Eppendorf Stream but Gilson, Sartorius and others exist.

A programmable repeat pipette that can accept a TXT file. The upload of a TXT file after calculating the amount of water to add to each well, and after specifying pipetting across rows or down columns, would allow the user to complete normalisation in as few as 9 pipetting steps: 8x12 channel 5ul transfers from the source plate to a normalisation plate, and 1x automatic repeat pipette dispense. "Simples" as the Meerkat says.

Nextera failures fixed by SPRI-cleanup

I'm continuing a series of posts that will highlight small advances we've made in the lab that are unlikely to be published in their own right, this time its the turn of SPRI-bead clean-ups and how we used them to rescue some samples that would not tagment.

The problem: We had a user submit a set of samples for Illumina's rapid exome sequencing in our library prep service. These samples were processed alongside others but the tagmentation for this one user was pretty awful. Most samples dropped out entirely and ony a few generated something that looked as if it had been tagmented at all. The figure below shows tagmentation before SPRI-clean-up and you can clearly see the control (inset) has tagmented and nothing is visible in the second lane where the users samples hold have been. You might also notice that the control is significantly under-tagmented we observed that our control DNA was too high in concentration (15ng/ul instead of 5ng/ul) hence the under tagmentation. Oops!

5mC-PCR: preserving methylation status during polymerase chain reaction

Methylation analysis is hampered by the simple fact that PCR amplification removes methylation marks from native DNA. We came up with a simple idea to produce a thermo-stable DNA methyltransferase to preserve methylation status through PCR cycles, allowing amplification of DNA and simplified analysis. The first thing we did was approach some enzyme companies to see if anyone had something suitable on their books, they did not. But they did seem to think this was a good idea so we designed a pretty simple experiment to test it: this involved going back to basics, to how PCR was performed before the application of Taq polymerase - we added Dnmt1 after each amplification cycle to copy methylation marks onto the daughter strands.

Hacking MiSeq updated and now hacking your BioAnalyser too!

In a post earlier this week I talked about the hacking of a MiSeq run by MadsAlbertsen, one comment on the post drew my attention to another paper I'd missed where the authors hacked their MiSeq to perform 600bp reads (PE300). Considering this was a year before Illumina sold us kits I'd say that's quite an achievement!

The Genome Sciences Centre, British Columbia Cancer Agency in Vancouver, did the sequencing for the Spruce genome paper (1). One of the authors is Robin Coope (Group Leader, Instrumentation BCCA Genome Sciences Centre) and he has been behind some pretty cool engineering in the genome sciences. In the Spruce paper his group demonstrated how to crack open a MiSeq cartridge and replace the insides with a larger reagent reservoir so kits can be mixed allowing much longer runs than Illumina intended (at the time of publishing).

The image below is from their supplementary data, I don't recommend you do this at home!

I met Robin when he was speaking at European Lab Automation 2013 in Hamburg last June. He gave an excellent talk on Automation Challenges in Next Generation Sequencing; we also had excellent weiner-schnitzel and dark bier once the conference finished. He spoke about the problems of quantifying NGS libraries on Bioanalyser and qPCR; we want molarity but get DNA concentration and these are not the same thing! Current methods allow you to use a simple calculation to convert between the two but this is heavily reliant on library size estimation. It is pretty much impossible to get the size right in the first place without measuring it and most people use the BioAnalyser. This is where Robin's talk really got interesting for me...
 
Unfortunately I can't share the slides (it was a commercial conference) but you could email Robin and ask him for a copy (or to hurry up and publish). Basically he described the deficiencies of the Bioanalyser software and introduced the concept of intelligent Fluorescent Units (iFU) to change the way the Bioanalyser does its analysis.

The Bioanalyser does a reasonable job of calculating size and molarity that works well on “tight” libraries, equally a visual estimation of mean insert size gives good results and cluster errors are more likely to be from mass quantification errors than insert size estimation errors. However for wider library distributions like Nextera or amplicons, iFU improves cluster density prediction and reduces cluster density error by 60% in the set (n=28) of amplicons he presented.

Of course none of this would be needed if we were using probe-based assays or digital PCR to count library molecules, but that is a whole other post!

Finally Robin went on to describe his groups work on the Barracuda a robot for 96 samples gel size selection.

Hack your MiSeq and get $400 off a 600bp run

I’ll start off by saying not quite, but you can read on to get an idea on how to increase read length of a MiSeq 500cycle v2 kit to get 600bp of data.

MadsAlbertsen posted on a SEQanswers thread about their protocol to squeeze a little more out of the MiSeq. They are using a hacked MiSeq Reagent kit v2 (500 cycles) and running a 2x301bp which is not supported by Illumina. “Do it at your own risk! (although it works nicely.)” is the message on the website. The group are using the modified protocol and hacked kits for bacterial 16S rRNA gene amplicon sequencing of the V1-3 variable region. The target region in E. coli position is a total of 489bp, but depending on target species can vary up to significantly (making the 2x301 run necessary).

How to hack your MiSeq kit: Make sure you follow the instructions on adding a little extra reagent to some of the wells.
5 mL of incorporation buffer from well 1 of a left-over reagent cartridge to well 1 of the 2x301 cartridge.
7 mL of scan mix from well 2 of a left-over reagent cartridge to well 2 of the 2x301 cartridge.
6.8 mL of cleavage mix from well 4 of a left-over reagent cartridge to well 4 of the 2x301 cartridge.
80 mL of incorporation buffer from a left-over incorporation buffer bottle.

Now simply set the Miseq to 2x301 in the samplesheet and ignore the warning the software gives. Et voila 600bp for the price of 500. With a MiSeq v3 kit costing about $1400 that’s potentially a $400 saving. 

Will we be doing this in my lab? No way, I’m far too conservative with users samples to play around like this. But I wish I could do more stuff like this, as it’s fun. It makes me want to come up with my own genomics Instructables.

Watch out for their paper: Saunders, A.M., Albertsen, M., McIllroy, S.J. & Nielsen, P.H. (in prep) MiDAS: the field guide to the activated sludge ecosystem.

Finding your way around NGS sample prep

I'm often asked which sample prep method a user should consider for their experiments. In my lab we use a lot of Illumina TruSeq kits; we've tried other methods, and do use Rubicon's Thruplex, but Ilumina's end-to-end support is useful in a medium sized core facility. And the kits work!

I wanted to illustrate the fact that Illumina sample preps share many steps in their protocols to demonstrate that once you've mastered one protocol, you can easily move onto another. The map of sample prep below is my first try at that illustration. You can see a higher resolution image here.

Explaining how the kits work always takes time and I have always thought a strength of the Illumina technology is the flexibility given by the core of sample prep: end repair, ligate adapters & PCR. Users can think creatively about what they do to their DNA (or cDNA) before starting or during the prep to come up with novel techniques. Stranded RNA-seq, bisulfite sequencing, RRBS, exome capture are just a few of the methods developed by users of Illumina technology. Hopefully the image above shows how similar things really are with those key steps clearly highlighted as large "interchanges", along with PCR and qPCR & Bioanlayser for QT/QC.

The above image borrows heavily from the London underground maps developed by Harry Beck in 1931. Most of the Illumina protocols are listed and I've included Thruplex on its own network for comparison. I'm aiming to add more detail for some of the different RNA-seq protocols at some point as well as bisulfite sequencing. I'm also thinking about how this might be extended to include details on suggested sequencing depth; lines for Human genome vs CNV-seq or DGE vs splicing.

Let me know what you think.

This work is licensed under a Creative Commons Attribution-ShareAlike 3.0 Unported License.

Targeted RNA-seq methods are here

Illumina and Life Technologies are both launching targeted RNA-seq applications which are likely to become standard tools for many labs; if the price is right.

The ability to target a portion of the genome has revolutionised next-generation sequencing experiments. The analysis of exomes has exploded, custom panels for exome-style pull-down are being used to great effect in 1000’s of samples and amplicon analysis is making it possible to run 10,000’s of samples in a single experiment (we’ve run a HiSeq flowcell with 12288 samples on it, 1536 per lane using Fluidigm – currently unpublished).

The next frontier looks like it could be using the same techniques to target a portion of the transcriptome, again allowing many 1000’s or 10,000’s of samples to be analysed in a single experiment. These technologies are likely to replace real-time PCR for mid- to high-plex studies. Anyone that has tried to run a few 100 TaqMan or SYBR assays on their 96-well qPCR machine will see the potential. And users with BioMark, TLDA, Wafergen, and other high-throughput qPCR systems will see the potential of using just one analysis method (NGS) as their primary data collection tool.

A better way to analysis of tumour heterogenity

Ben Raphael's lab at Brown University have just published THetA for tumour heterogeneity analysis; it's a great paper so download it and have a look yourselves.

Solid tumours have been shown to be highly heterogeneous and that this can underlie drug resistance and eventual relapse in patients. Individual tumours are admixtures of different tumour-cell sub-populations and NGS data come from this mix. The THetA paper introduces a method to infer the starting population of distinct tumour-cells.

5hmC on Illumina 450k's: analysis without breaking the bank?

Epigenetics is one of the most exciting areas of research and promises to impact all sorts of biological systems. DNA methylation has recently been shown to come in methylcytosine (mC) and 5-hydroxymethylcytosine (5hmC) flavours. Usually analysis of mC is carried out using bisulfite conversion or enzymatic digestion. The gold standard is probably bisulfite sequncing and a few groups have made significant advances using Bis-seq methods on Human genomes.

Moleculo data presented at Illumina Scientific Summit

Geoff Smith from Illumina presented a great piece of work his group have been collaborating on with Jill Banfield’s research group at the University of California Berkely, Department of Earth and Planetary sciences. Geoff included a discussion on their use of Moleculo.

NGS forensics might require unique barcodes

The use of NGS in forensics is likely to supplant current STR profiling. One issue that has been discussed is the need to robustly determine that the correct sample has been analysed and reported on. With NGS almost all sequencing today is being done in a multiplex fashion using indexing barcode reads of generally around 6-10bp.

Current barcoding uses sets of up to 96 or 384 barcodes which is fine for many research applications. However there are reports of carry-over of barcodes from one run to another  on the kinds of instruments that are likely to be used in forensic labs. This is likely to make courts nervous about bringing this technology in.

One way to remove some of the problems would be to create significantly longer index oligos that are unique. I can see a market for a consumable kit where each is truly unique. The indexes included would be single-use only and no-one else in the world would get the same index oligo. Oligos can be made in very small synthesis scales today and in high-throughput. Given a long enough read even lower quality synthesis should allow confident discrimination of contamination allowing users to be confident about the final results.

A 25bp barcode read should be as unique as we might ever need and on HiSeq today would not add much time to a sequencing run.

Using SPRI beads to improve Nextera

We have been running Nextera in my lab for a while now and had some great success e.g. 600x C. elegans genomes. With the release of Illumina's rapid exome kits Nextera is proving to be a versatile and useful addition to our lab tools. Unfortunately it has at least one significant weakness; the need to accurately quantify dilute DNA.

Automating Illumina library preparation

Illumina's just announced a partnership with major automation vendors so their s-bot looks like it has been killed as a project. We've got a Tecan Evo in our lab and would like to use it for library prep but a major headache has been the rapid development of library prep methods. There just does not seem time to program and validate a method before the next one comes out.

2D RNA-seq rocks!

RNA-seq is getting very cool over in Sweden's SciLifeLab. A recent press release announces that Joakim Lundeberg (Royal Institute of Technology) Jonas Frisén and Patrik Ståhl (Karolinska Institutet) were awarded £1.8M from the Knut and Alice Wallenberg Foundation to study Brain biology. They have developed a method of 2-dimensional gene expression, 2D RNA-seq and will use this to comprehensively map gene expression in the brain. They also aim to compare normal and diseased brains, including cancer, autism and schizophrenia.

ENCODE's RNA-seq recommendations need revising

One of the great things about ENCODE was the amount of effort put into standardising how different groups did their experiments. As any core manager knows a good SOP is a huge step to high-quality data. The ENCODE consortium put together guidelines for ChIP, RNA and RIP. Those guidelines, published almost a year ago, are due for revision in a month or so. I'm looking forward to the revised document as the current recommendations have some serious problems from my personal experience.

ctExome-seq: Circulating tumour exomes as a non-invasive method to track tumour evolution during treatment

Monitoring of cancer patients is an important part of their treatment and this is traditionally done with tests like Computed Tomography imaging (CT) or biomarker analysis. NGS of cancer amplicons, exomes and/or genomes is being discussed as a realistic addition, and possibly an alternative, to these traditional methods. A major hurdle to their use is the fact that analysis of cancers can be hugely complicated by tumour heterogeneity and that serial biopsy of patients to obtain material for sequencing is not a realistic option for many patients. Although the analysis of circulating tumour cells has received a lot of interest, collecting the cells is no easy task (there have been some great papers that show what might be possible from CTC’s, the MALBAC method published in Science is a current favourite of mine). Nitzan Rosenfelds group at the Cambridge Institute has been pioneering the use of circulating tumour DNA (ctDNA) sequencing as a more amenable method to understand cancer genomics.

Mate-Pair made possible?

Mate-Pair sequencing has been used from the early days of genome sequencing to help with final assembly. The technique creates libraries with much larger inserts than the standard fragment library prep, even 10Kb or more. However making these libraries has never been easy and the methods used have not changed a great deal from the days of the HGP.

A few groups have been successful in creating mate-pair libraries but my experience has not been so good. My lab has only tried once and did not get great results, many of the groups I work with have found the preps difficult and sequencing results less than encouraging. It looks like mate-pair is a technique where “green fingers” are required. This is not a situation I like as I am a firm believer that as long as a protocol has been carefully put together then anyone should be able to follow it. 

How does mate-pair work: High-molecular weight genomic DNA is fragmented to an approximate size range, usually 3, 5 or 10Kb, this is then end-repaired with biotinylated nucleotides and gel-size selected more specifically. Fragments are then circularized by ligation, and purified by biotin:streptavidin cleanup. These circularized fragments are then subjected to a second round of fragmentation (to 300-500bp) biotin:streptavidin cleanup to remove all but the ligated ends of the original molecules. This DNA is then used for a standard fragment library prep and mate-pair sequencing. The length of the initial gel size-selection determines the mate-pair gap size expected during alignment of the final sequence reads. The orientation of these reads also acts as a useful QC. Two major problems are the amount of DNA required, usually 10ug or more and the creation of chimeric molecules during ligation that produce artifactual structural variations. 

Jan Korbel’s group at EMBL have been successfully using mate-pair sequencing for their analysis of structural variation in Cancer (Paired-End Mapping Reveals Extensive Structural Variation in the Human Genome). They have been using a pretty standard protocol. They fragment DNA using a Hydroshear (see the bottom of this post for more details), cut gels for 5Kb libraries, and try to minimise PCR to keep diversity high. They are processing 8 samples at a time and it takes 5 days but the tedious and careful handling pays off in some very nice results. The biggest drawback is the requirement for 10ug of DNA.

Newer methods offer some promise for simpler mate-pair prep: At least two new products have been released that may help the rest of us produce reliable mate-pair libraries. Lucigen have developed a new BAC/Fosmid vector system for the creation of high-quality libraries with large inserts. Illumina have released a new Nextera based mate-pair kit. 

Lucigen pNGS: Lucigen's novel protocol for making clone-free libraries of 40-300kb insert size could have a dramatic impact on the de novo assembly of complex genomes. They have released the pNGS system that includes a di-tagged vector which contains the PCR amplification sites required to produce sequence ready libraries. This vector lacks the promoter for lacZ and has transcriptional terminators which, according to Lucigen, “result in higher stability cloned insterts”.

Lucigen workflow

The system works by using random DNA fragmentation to produce inserts of 40-300kb that are gel-purified and cloned into the pNGS vector. The amplified BACs or Fosmids are then digested with 4bp cutters to leave just the ends of the original insert fragment and the vector sequence. This digested molecule is self-ligated to create a circular template for NGS PCR amplification producing the final sequencing library. 

Nextera mate-pair: The new protocol from Illumina makes use of the Nextera product Illumina acquired when they bought Epicentre in 2010. The protocol is not completely what I expected as it uses a mix of Nextera and TruSeq reagents rather than relying on Nextera alone. Illumina have a TechNote on their website technote_nextera_matepair_data_processing that I'd recommend interested readers take a look at. They show comparison data from paired-end vs mate-pair+paired-end sequencing of the Human genome and show a modest, but important increase in coverage statistics that is most obvious in repeat regions of the Human genome.

Nextera mate-pair offers gel-free or gel-based size selection. The gel-free option allows a lower DNA input of just 1ug but generates a larger final mate-pair size distribution of 2-15kb, which may make analysis harder as you cannot simply discard mate-pairs using insert size as a QC. The gel-based protocol requires 4ug of DNA but the user has control over the final mate-pair size distribution as normal.

Transposomes are loaded with biotinylated oligs and are mixed with DNA at a much lower ratio than in the standard Nextera kits, which allows much larger fragment sizes to be produced (2-15kb). The mate-pair library is then size-selected and circularized as in standard protocols (this is still a blunt-ended ligation which is less efficient than the “sticky” one used in the Lucigen kit. Physical shearing breaks up the large circular molecules and the biotinylated oligos added by in vitro transposition allow capture of only the mate-pair regions. These are purified and used as the template to a standard TruSeq library prep.

I like Nextera and we have been using it in capture projects and the new XT formulation. My ideas about what Nextera mate-pair might look like made use of two sets of transoposome loaded with different sequences. The first would create the large fragments, incorporate biotin and leave compatible ends for a "sticky" ligation. The second would be a standard Nextera prep to produce the final library which could be streptavidin purified before PCR.

Will structural-variants in Cancer be easier to detect: We’ve done very little mate-pair in my lab because of the sample requirements, so I'm hoping that these new developments will mean more users request the protocol and are able to make use of the additional structural variation data. For now many people seem to be happy with getting 80% or more of the variants from standard fragment libraries. However protocols that allow generation of multiple mate-pair sizes that can be indexed for sequencing are likely to allow identification of important, and so far difficult to identify rearrangements in Cancer genomes. Being able to run a single pooled sample that contains tumour:normal at ~350, ~3Kb, ~10kb & ~40kb inserts should give very high resolution copy number and high-quality structural variation data. This may also be achievable with far fewer reads than are used today and with the bonus that significantly less DNA is used in the prep.

Hydroshear vs Covaris vs Soniction vs enzymes: There are many ways to chop DNA into fragments but only a few will reliably give the larger ones required for successful mate-pair library preparation. Most of us are using Covaris or Bioruptor to produce 300-500bp fragments. These instruments can also generate longer bits of DNA but they are inefficient as most DNA is outside the range required.

The Hydroshear is a very clever piece of kit from Digilab that uses a pretty simple mechanism to break DNA into relatively tight fragment distributions. DNA is pushed through a tight contraction in a tube by a syringe. As the sample moves through the contraction the flow rate increases dramatically, this stretches the DNA until it snaps. The process is repeated over several cycles until an equilibrium is reached. The flow-rate and the size of the contraction determine the final fragment size. The smear of DNA usually seen on a gel after sonication is much tighter, e.g. 1.5-3Kb.

I'm writing a post on DNA fragmentation methods and will go into more detail there.

How do SPRI beads work?

Someone recently asked me, “how do SPRI beads work” and I realized I was not completely sure so I went to find out.

My lab uses kits. Lots and lots of kits; kits for DNA extraction, kits for PCR, kits for NGS library prep and kits for sequencing. Kits rock! But understanding what is going on at each stage of the protocol provided with the kit really helps with troubleshooting and modification. How many people have added 24ml of 100% Ethanol to a bottle of Qiagen’s PE buffer without stopping to ask what is already in the bottle? The contents of the PE bottle are… answer at the bottom of this post.

I hope this post helps you understand SPRI a bit better and think of novel ways to use it. I’d recommend reading A scalable, fully automated process for construction of sequence-ready human exome targeted capture libraries in Genome Biology 2011 for the with-bead methods discussed at the bottom of this post. I am sure we'll all be using their method a lot more in the future!

You are most likely to come across SPRI beads labeled as Ampure XP from Beckman.

First some handling tips:

• Vortex beads before use.
• Store beads at the correct temperature.
• Allow enough time for beads to come to room temperature.
• Pipetting is critical; use very careful procedures (SOPs) and well calibrated pipettes.

How do SPRI beads work? Solid Phase Reversible Immobilisation beads were developed at the Whitehead Institute (DeAngelis et al 1995) for purification of PCR amplified colonies in the DNA sequencing group. SPRI beads are paramagnetic (magnetic only in a magnetic field) and this prevents them from clumping and falling out of solution. Each bead is made of polystyrene surrounded by a layer of magnetite, which is coated with carboxyl molecules. It is these that reversibly bind DNA in the presence of the “crowding agent” polyethylene glycol (PEG) and salt (20% PEG, 2.5M NaCl is the magic mix). PEG causes the negatively-charged DNA to bind with the carboxyl groups on the bead surface. As the immobilization is dependent on the concentration of PEG and salt in the reaction, the volumetric ratio of beads to DNA is critical.

SPRI is great for low concentration DNA cleanup that is why it is used in so many kits. The reagents are easy to handle and a user can process 96 samples very easily in a standard plate. Alternatively the protocol can be easily automated and tens or hundreds of plates can be run on a robot in a working day. The binding capacity of SPRI beads is huge. 1ul of AmpureXP will bind over 3ug DNA.

This is the typical SPRI protocol from Beckmans website.

Size-selection with SPRI: Again the concentration of PEG in the solution is critical in size-selection protocols so it can help to increase the volume of DNA you are working with by adding 10mM Tris-HCL pH 8 buffer or H2O to make the pipetting easier. The size of fragments eluted from the beads (or that bind in the first place) is determined by the concentration of PEG, and this in turn is determined by the mix of DNA and beads. A 50ul DNA sample plus 50ul of beads will give a SPRI:DNA ratio of 1, as will 5ul pipetting (but much harder to get right). As this ratio is changed the length of fragments binding and/or left in solution also changes, the lower the ratio of SPRI:DNA the higher larger the final fragments will be at elution. Smaller fragments are retained in the buffer usually discarded and so you can get different size ranges from a single sample with multiple purifications. Part of the reason for this effect is that DNA fragment size affects the total charge per molecule with larger DNAs having larger charges; this promotes their electrostatic interaction with the beads and displaces smaller DNA fragments

SPRI size selection from Broad "boot camp"

SPRI size selection from Broad "boot camp"

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Broad Boot Camp http://www.broadinstitute.org/scientific-community/science/platforms/genome-sequencing/broadillumina-genome-analyzer-boot-camp


With-bead SPRI cleanup: An incredibly neat modification of SPRI clean-up is the “with-bead” method developed by Fisher et al in the 2011 Genome Biology paper. Rather than using SPRI to clean-up discrete steps in a protocol these are integrated into a single reaction tube method, thus reducing the number of liquid transfer steps. After each step DNA is bound to beads by addition of the 20% PEG, 2.5M NaCl buffer, washes are performed as normal with 70% ethanol but the DNA is not eluted and transferred. Rather the master-mix for the next step in the protocol is added directly to the tube. The final DNA product is eluted from beads for further processing e.g. Illumina sequencing. This modification increases DNA yields in Illumina library prep as multiple transfer steps are removed, reducing the amount of DNA lost at each transfer.


How will you use SPRI in your lab?

PS: the Qiagen PE bottle contains 6ml of water.

PPS: If DNA and Carboxyl are both negatively charged what is happening on a SPRI bead? There appears to be a bit of a black hole in the literature I read through about how the SPRI beads actually work at the molecular level. Digging around some more it appears that the PEG can induce a coil-to-globule state change in the DNA with the NaCl helping to reduce the dielectric constant of the solvent, PEG also acts as a charge-shield. These mechanisms may be behind the "crowding" talked about on other sites. Perhaps someone out there can enlighten me?


Refs:
Lis JT. Size fractionation of double-stranded DNA by precipitation with polyethylene glycol Methds Ezymol (1975)
Lis J. Fractionation of DNA fragments by polyethylene glycol induced precipitation. Methds Ezymol (1980)
DeAngelis et al. Solid-phase reversible immobilization for the isolation of PCR products. NAR (1995)
Hawkins et al. DNA purification and isolation using a solid-phase. NAR (1994)

Can gene expression be considered a quantum problem?

I had a very interesting discussion over coffee yesterday at TRON, a new Translational Oncology institute in Mainz, Germany. TRON is specialising in cancer immunology and genomics, or Immunomics. One of the things they are aiming to do is develop personalised cancer vaccines by finding imunogenic markers in tumours and using long RNA molecules to present these markers to T cells to activate and modulate the imune respsones.

I was giving a seminar about technology development in next-generation sequencing, during the day I spoke with several people about single-cell analysis.

A problem with analysing a single cell is you only get one chance to take a look at its molecular biology as we use destructive methods to assay the cells genome and transcriptome. Single-cell gene expression usually requires the use of RNA amplification methods with their own biases and this led the discussions to a question.

Is RNA expression limited by quantum mechanics and is their a fundamental limit on the accuracy with which the level of gene expression can be measured in its true biological context? Analysing a single cell means analysing the whole transcriptome as a snap-shot of a dynamic biological system. Our discussions went around the question "is the absolute measurement of any one gene to some extent meaningless, when asking questions about what that level of gene expression means in terms of biology for that cell?"

We discussed technical issues with single-cell gene expression analysis and both TRON and CRI are using the Fluidigm gene expression chips for this kind of analysis. However replicate numbers need to be high and analysis methods probably need to take into account variance in gene expression as much as mean expression levels and fold-changes.

What is "total RNA"?

I wrote this post after a recent experimental design discussion where we went round and round discussing RNA analysis.

There are many different RNA molecules in a cell. When I was studying there were only three, mRNA, ribosomal RNA and tRNA. Simple! However over the last couple of decades our understanding of the complexity of RNA transcription and what different RNA's do in the cell has changed dramatically. Regulatory RNAs have become incredibly important to consider in biological systems and include both small RNAs including miRNA, piRNA and long-noncoding RNAs such as Hotair. These can target mRNA for degradation, block mRNA translation or even target genes for methylation and block transposon mobility.

We are still finding out an awful lot about the transcriptome that is surprising, and controversial (e.g. RNA editing).

Unfortunately our earlier ignorance of the varied RNA landscape has meant the term ‘total RNA’ is somewhat confused. As more and more scientists look deeper into the transcriptome and RNA biology, it is important to understand what total RNA is a particular experimental context. I try to understand what it is the experimenter wants to assay and use that as the starting point for discussions about particular extraction chemistry, labelling and other methodologies.

Just for the record, for me total RNA means all the RNA in a cell.

What is in Total RNA: It’s all the RNA in a cell and includes lots of different RNAs and certainly not just messenger RNAs!

Here I have suggested a pretty comprehensive list but feel free to point out any I may have missed: mRNA, polyA RNA, polysomal RNA, tRNA, ribosomal RNA, lincRNA, miRNA, piRNA, siRNA, SRP RNA, tmRNA, snRNA, snoRNA, SmY RNA, scaRNA, gRNA, aRNA, crRNA, tasiRNA, rasiRNA, 7SK RNA

A historical perspective and Affymetrix U95A arrays: When I first started running gene expression arrays in 2001 we often saw degraded RNA at the bottom of a gel or Bioanalyser trace. Many users wanted to clean up this degraded RNA and so we would run a column based clean-up which effectively removed all the small RNA species from a sample. Nearly all extractions were performed with Trizol or something similar and column-based extractions were the exception rather than the rule.

Back then we were only interested in gene expression assayed by labelling mRNAs by oligo-dT primed cDNA synthesis in an Eberwine amplification reaction. We were not aware that in that degraded RNA we were so happy to remove was an undiscovered regulatory universe of RNA molecules. The techniques we were using to assay gene expression would not have picked these up anyway, but nether-the-less we made a decision based on a very naive understanding of RNA biology. We should all have learnt from that lesson.

RNA extraction: Now when asked for a recommendation on RNA extraction I usually suggest Qiagen miRNeasy or another kit that preserves the small RNA fraction in a sample. Some users come to us with RNA already prepared and ask if we can also look at miRNAs but if they have used a kit that does not keep these small molecules we have to suggest they start again from scratch.

Kits that preserve small RNAs are only a little more expensive to use than others, and they do not involve much more lab work. Even if someone is interested in mRNA gene expression I always try to persuade him or her to collect small RNA containing total RNA. For many experiments the samples will only be collected once and might be reused in experiments many months or even years later.

What about assay biochemistry: Even if you have collected total RNA it is not necessarily possible to analyse all the different types of transcript in a single assay. Most mRNA gene expression arrays use very different labelling methods than microRNA arrays for instance. You can’t very easily label mRNAs and miRNA with the same chemistry.

With Illumina and other RNA-seq methods most users need to choose whether to assay mRNAs or micro-RNAs and use different methods to do so that use very different biochemistry. Whilst it is possible to use an RNA ligation method to analyse small RNAs and fragmented mRNAs very few people do so as this is more complex to prepare and analyse than running two independent experiments with the same samples as inputs.

What should you do for your next RNA experiment: If you are about to run a microarray or an RNA-seq experiment I’d encourage you to take a look at some of the literature out there and read the manuals for several kits to understand what is happening to RNA molecules in a particular method. By doing this you will be better placed to decide how best to approach your experiment and understand the limitations of the method you have chosen. An excellent resource is the Ambion website.

If you are lucky enough to have a good core-lab or service provider to run your experiment then take the opportunity to talk to them before you start the experiment. This means before you get cells out of the freezer or tissue from your mice, and hopefully you can work together on an experimental design that will get you the most from the samples. The extraction may only cost you a few £, $ or ¥ extra and make your day an hour or two longer in the lab, but you’ll be happy you did it when someone publishes something interesting about the regulation of your favourite gene in a similar sample set, as you’ll be able to go back to the freezer and confirm their finding in the samples you already have.