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Old 11-13-2014, 12:08 PM   #1
esherman
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Default 'Assembly PCR' for homemade amplicon library prep

I'm trying to make homemade amplicon library preps for the Illumina MiSeq, and my fusion primers always end up being around 100bp (if not more!) by the time I add all the necessary sequences (p5/p7, indices, primer landing sites, N's for diversity, etc.). This requires I buy oligos at the 250nm scale of synthesis rather than the standard 25nm scale. When I'm buying a whole plate of 96 indices, this gets very expensive very fast!

I'm thinking of an 'assembly PCR' approach in which I order four 25nm oligos. Two short ones that contain the sequence-specific sequence fused to the Illumina sequencing primer landing sites and two additional short ones that contain the p5/p7 sequences, indices, and a bit of the primer landing site. These could all be thrown into a single PCR reaction (perhaps with a bias towards the 'outer' primers containing the p5/p7 sequence).

Of course I would have to pay attention to strandedness and directionality when designing the oligos, but I wanted to see if anyone has tried this before I spend too much time thinking about it.

This idea would also have the benefit of being able to change out the sequence-specific 'inner' primer and re-use the outer primer/indices for different experiments.
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Old 11-13-2014, 02:37 PM   #2
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I think Illumina recommends a two-step approach. Fusion primers with barcode and sequencing primer locations, then a second PCR with P5/P7 ends being added. If you do it all at once it may get ugly. Also if you are low diversity adding in a N+1, N+2 linker in there greatly helps diversity.
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Old 11-13-2014, 02:48 PM   #3
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Try this:

http://supportres.illumina.com/docum...15044223-b.pdf

Also, I'm not sure you need the extra N bases to increase diversity, if you're using the newest MiSeq software.

Cheers,

Scott.
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Old 11-13-2014, 05:33 PM   #4
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Hi esherman,

A few options/opinions to offer on this.
1) If you do decide to go 2-step you risk contamination. These workflows are really really hard to keep clean. Personally I 'hate' this approach but it does kind of depend on what you are using the data for.
2) Do you need to do paired end sequencing? - we are getting good single direction sequencing out to 250+ bases (on v2 300 nano kits), so if your amplicon is shorter than that you could consider it (i.e you only need a sequencing primer at one end).
3) Use a custom sequencing primer (e.g one with LNA's) to shorten the the total length of the primer you need. Search some of the seq-answer threads on this for a few hints or PM me.
4) a combination of points 2+3 above is what we (mostly) use to keep primer sizes down which helps cost and PCR efficiency.

best of luck with your workflows, Cheers Mike

Last edited by bunce; 11-13-2014 at 06:16 PM.
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Old 11-13-2014, 05:54 PM   #5
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I'd second the two-step approach. Do a first round with a fusion primer with the specific sequence and full/partial read primer sequence followed by a second round that adds the indexes and p5/p7.

That way gives you the most flexibility, as well, because you can keep reusing the indexing primers for future experiments and can always switch around which library gets which index so you don't end up with incompatible overlap.

As for cross contamination, having separate pre and post PCR benches is critical (much more common in corporate labs), but you shouldn't need to do many PCR cycles at all to add the p5/p7 and indexes, so you really just need to make sure your first PCR product and final libraries stay the hell away from the bench where you set up the first round of PCR.
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Old 11-13-2014, 06:30 PM   #6
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Quote:
Originally Posted by cmbetts View Post
As for cross contamination, having separate pre and post PCR benches is critical (much more common in corporate labs), but you shouldn't need to do many PCR cycles at all to add the p5/p7 and indexes, so you really just need to make sure your first PCR product and final libraries stay the hell away from the bench where you set up the first round of PCR.
I would not be so hasty in talking down the contamination risk in amplicon workflows - our ancient DNA set-up is 'extreme' and we still get bleed-though of tags (we never re-use index pairs). If you are doing a 2nd round PCR you still need to open that tube in a post-PCR 'area' to spike in the 1st round PCR. Amplifying amplified DNA that then undergoes another amplification (cluster generation) can cause headaches downstream from chimera formation to contamination that builds over time. But, again, it does depend on the end use of the data. thats my 2cents worth - cheers, Mike
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Old 11-13-2014, 08:44 PM   #7
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As a rule of thumb, we never use the same barcode combination twice on a plate. It's not like you have the option of running the same barcodes in multiple lanes like you can on a HiSeq.

For us the most critical part is keeping your primer stocks contamination free prior to the PCR. After normalization and pooling you would have no way of knowing your DNA was tagged with multiple barcodes without some sort of internal control or verification of the target.

Pipette wisely!

Last edited by DNA_Dan; 11-13-2014 at 08:49 PM.
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Old 11-14-2014, 12:25 AM   #8
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Quote:
Originally Posted by bunce View Post
I would not be so hasty in talking down the contamination risk in amplicon workflows - our ancient DNA set-up is 'extreme' and we still get bleed-though of tags (we never re-use index pairs).
I wonder if you have considered possibility of carry over from a previous run on MiSeq. We use two step PCR and it works well. In addition, it is convenient and we can pool and sequence other libraries as well without risk of unwanted interaction between custom and Illumina sequencing primers. Nano is the most expensive option for Illumina systems per Gb of data.
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Old 11-15-2014, 11:53 PM   #9
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Quote:
Originally Posted by nucacidhunter View Post
I wonder if you have considered possibility of carry over from a previous run on MiSeq. We use two step PCR and it works well. In addition, it is convenient and we can pool and sequence other libraries as well without risk of unwanted interaction between custom and Illumina sequencing primers. Nano is the most expensive option for Illumina systems per Gb of data.
Hi, Sure - carry-over is always a possibility (we have been using the bleach wash protocol since february). I suspect (with no evidence) that people may 'blame' carry-over when it is on fact the lab workflow causing issues.

I don't doubt that a 2-step PCR protocol works well - it will always generate a truck-load of library. The question is what portion or reads are chimeric or have tags that you have not used on that run? I would be interested if anyone (using 2-step PCR and re-using indexes), has quantified this?
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Old 11-16-2014, 11:57 PM   #10
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Quote:
Originally Posted by bunce View Post
Hi, Sure - carry-over is always a possibility (we have been using the bleach wash protocol since february). I suspect (with no evidence) that people may 'blame' carry-over when it is on fact the lab workflow causing issues.
Illumina has acknowledged carry over issues: https://icom.illumina.com/MyIllumina...applications-m

If by “evidence” you mean how many people look for evidence of contamination source in their reads, I do not have an answer.

Quote:
I don't doubt that a 2-step PCR protocol works well - it will always generate a truck-load of library. The question is what portion or reads are chimeric or have tags that you have not used on that run? I would be interested if anyone (using 2-step PCR and re-using indexes), has quantified this?
Two step PCR if done properly does not generate truck load of amplicons. The aim should be number of cycles to generate enough yields for a QC and 3-4 sequencing runs which usually would be 50 ng>.

Chimeric reads actually would be more prevalent in one step PCR rather than two step, because two major reasons for Chimera formation during PCR are incomplete extension and strand invasion. The condition for both are favoured in one step PCR where the reagents are most likely to deplete and concentration of amplicons increase to a critical point favouring those reactions.

By using indexed primers for second step from a plate sealed with a plastic film and frozen, thawed and resealed multiple times, I have got 1 in 10,000 reads that had indices not used in the reactions. By using individual tubes containing an indexed primer and opening them one at a time such as recommended in Nextera protocol, no read was detected that had unused indexed primer. One obvious advantage using two step PCR with dual indexing (such as one described in Illumina 16S sequencing protocol, see link in #3) is that identifying and eliminating cross contamination post-sequencing (caused either by physical contamination during library prep or image analysis error with higher cluster densities) would be easy as chance of both primers being contaminated is reduced.

Quote:
Amplifying amplified DNA that then undergoes another amplification (cluster generation) can cause headaches downstream from chimera formation to contamination that builds over time. But, again, it does depend on the end use of the data.
Cluster generation should not encourage chimera formation because bridge amplification takes place in solid state and unlike in solution PCR, fragments are not free to interact with each other. Fragments that are in close proximity or hybridize to each other and may fuse and form a cluster, will not produce pure signal either because of presence of mix template amplification or multiple primer binding sites and will fail filter so they cannot contribute to chimera reads.

It is not clear to me what sort of contamination would built over time, if one follows standard molecular biology lab hygiene such as using disposable gloves, aerosol free tips and cleaning bench and pipette surfaces.
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Old 11-17-2014, 12:52 AM   #11
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Hi nucacidhunter, I guess we will agree to disagree on some of these points. I can't conceive of a way in which a 2-step protocol can be 'cleaner' or less susceptible to artefacts than a 1-step set-up. It may be cheaper, arguably more efficient? - but not cleaner. When it comes to NGS, PCR is a necessary evil it should be minimised in anyway possible.

Quote:
Originally Posted by nucacidhunter View Post
Chimeric reads actually would be more prevalent in one step PCR rather than two step, because two major reasons for Chimera formation during PCR are incomplete extension and strand invasion. The condition for both are favoured in one step PCR where the reagents are most likely to deplete and concentration of amplicons increase to a critical point favouring those reactions.
Your logic is lost on me here as the 2-step method also goes though a 1st round PCR that is just is susceptible to Chimeras (I think we agree that PCR cycles should be kept to a minimum). In amplicon sequencing some people work indexes into their 1st round PCRs then pool and amplify (as a pool) to get p5-rd1/rd2-p7 onto the products. In these situations there has been a number of reports of 'jumping' indexes presumably the result of incomplete extension. This "can't" happen (i.e. index jumping) in a 1-step workflow as the forward and reverse indexes are the only ones in tube.

Quote:
Originally Posted by nucacidhunter View Post
By using indexed primers for second step from a plate sealed with a plastic film and frozen, thawed and resealed multiple times, I have got 1 in 10,000 reads that had indices not used in the reactions. By using individual tubes containing an indexed primer and opening them one at a time such as recommended in Nextera protocol, no read was detected that had unused indexed primer.
If a 1/10,000 contamination rate suits your application then that is good - it won't suit everyone. People not as adept at removing those pesky films may report a higher rate????

Quote:
Originally Posted by nucacidhunter View Post
One obvious advantage using two step PCR with dual indexing (such as one described in Illumina 16S sequencing protocol, see link in #3) is that identifying and eliminating cross contamination post-sequencing (caused either by physical contamination during library prep or image analysis error with higher cluster densities) would be easy as chance of both primers being contaminated is reduced.
This is is a way to identify contamination but is not an "obvious advantage " over a 1-step library generation. A 1-step workflow that integrates indexes and adapters at the 1st PCR so contamination is just as easy to spot.


Quote:
Originally Posted by nucacidhunter View Post
Cluster generation should not encourage chimera formation because bridge amplification takes place in solid state and unlike in solution PCR, fragments are not free to interact with each other. Fragments that are in close proximity or hybridize to each other and may fuse and form a cluster, will not produce pure signal either because of presence of mix template amplification or multiple primer binding sites and will fail filter so they cannot contribute to chimera reads.
Agreed, chimera's are not really an issue at cluster stage. But it is an amplification stage so one contaminatiing template molecule could initiate that cluster.


Quote:
Originally Posted by nucacidhunter View Post
It is not clear to me what sort of contamination would built over time, if one follows standard molecular biology lab hygiene such as using disposable gloves, aerosol free tips and cleaning bench and pipette surfaces.
.

Aerosols build in labs over time. In a post PCR area you can use ART tips, UV and gloves but this minimises contamination it won't remove it. A good PCR once 'opened' will generate aerosols with 10^5-10^9 copies of DNA that can travel through HEPA filters and build in a lab.

But to bring this banter back to esherman's question - there are good ways to generate amplicon libraries using both 1-step and 2-step methods there are strengths and drawbacks to both approaches. How you tackle this is very much dependent on budget, sensitivity, contamination concerns and the application you intend for the data.
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Old 11-17-2014, 02:14 AM   #12
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Hi bunce, I am fine with disagreement as far as it is evidence based and I agree with you in that cleanliness or tolerance of contamination is dependent on application and other factors. In two-step PCR usually first PCR goes for 15-20 cycles and then an aliquot is used for second PCR, say for another 10 cycles. This would generate fewer artefacts than a similar one-step PCR cycled 30 times because the amplicons would have less concentration and PCR reagents are replenished reducing the chances of incomplete extension and high concentration of amplicons two major cause of PCR artefacts.

The level of cleanliness you are hinting is more relevant to forensic and ancient DNA work which unfortunately most of the time suffers from contamination during or even before sample collection.

Out of interest is there any evidence that how many of those 10^5-10^9 aerosol copies of DNA that travel through HEPA filters and build in a lab, contaminate the work being done in that lab.
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Old 11-19-2014, 09:09 AM   #13
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Seems to me that one of the biggest 'deterrents' of contamination between samples within one prep, between preps across time, etc is to use enough template DNA so that any contamination won't be preferentially amplified.

Our platform may be more-ammenable to this type of prep, as we know our distal ends of the DNA to be sequenced…

We use a 2-stage PCR to build out our Illumina ends. As suggested earlier, the primary PCR uses library-specific oligos, but with about 1/2 of the Illumina adapters extending off the ends.
The Secondary PCR primers introduce the P5/P7 ends, as well as the barcode, but have no complementarity to the initial library, only the distal ends of the primary PCR product that are part of the illumina adapter just inbound of the barcodes.

We perform a PCR clean-up between each step to clean out the primers…
We use 10ng of input DNA into a 50ul PCR reaction, and that amount of template is a ton more than any contamination event unless you're an idiot...
Also, in-line PCR negative controls are always run…
And we don't ever sequence with the same barcodes two runs in a row...
And I am planning on installing the Bleach wash, but we typically only see about 1% of the total yield of DNA going into the 'undetermined' fastQ file.

I got fancy once, and added 1/1000th the amount of primary PCR primers for 2 cycles, and then added in the typical amount of secondary primer (barcode, P5P7)…that works but I don't like opening fresh PCR products and adding more primers to do more PCR. now that's asking for contamination.
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Old 11-19-2014, 05:51 PM   #14
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Quote:
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Seems to me that one of the biggest 'deterrents' of contamination between samples within one prep, between preps across time, etc [U]is to use enough template DNA so that any contamination won't be preferentially amplified
.

I agree - it is a numbers game. The issue however is that some applications are setup to find rare variants (e.g. circulating cancer cells or low concentrations of microbes in a complex mixture) - in these scenarios a low level (and unpredictable) baseline of contamination can be problematic. Again, (to sound like a broken record) the workflow - be it 1-step and 2-step - depends on end-use of the data.

The point I am trying to make here is that contamination in NGS workflows is real and needs to be managed - never more so than when amplifying amplified DNA.
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Old 11-20-2014, 10:24 AM   #15
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Contamination aside, have any of you looked at the effect of resampling bias from doing so much PCR on a sample? I have a customer looking to do this approach coming from RNA. This means 4 rounds of PCR, 1 RT reaction, 1 target specific nest, then the Illumina 2 step. I plan on titrating yield on a Lonza gel to optimize PCR cycles, but still sounds like a lot of resampling.
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Old 11-22-2014, 12:05 AM   #16
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Aerosols build in labs over time. In a post PCR area you can use ART tips, UV and gloves but this minimises contamination it won't remove it. A good PCR once 'opened' will generate aerosols with 10^5-10^9 copies of DNA that can travel through HEPA filters and build in a lab.
If I understand you correctly, you are saying that contamination by PCR is inevitable and nothing can be done about it. I have to disagree with you because PCR has been and is and probably will be corner stone of molecular biology.

If I open my PCR tube in a PCR cabinet with outside filtered air pushing through, any aerosol will be pushed away by air flow. Any possible remaining amplicon can be degraded by UV light or cleaned chemically and cabinet will be free from any amplicon. The question is what evidence you have or you can refer to show that usual lab hygiene is not sufficient to prevent PCR contamination. An amplicon that supposedly sticks to lab wall, how likely can be airborne again and land in a PCR tube. In a very slim probability if it lands how that really is going to affect results because the target amplicons will be in billions.

I strongly, would suggest external expert review of ancient DNA set up and practices if they can not control PCR contamination.
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Old 11-22-2014, 12:41 AM   #17
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If I understand you correctly, you are saying that contamination by PCR is inevitable and nothing can be done about it. I have to disagree with you because PCR has been and is and probably will be corner stone of molecular biology.

If I open my PCR tube in a PCR cabinet with outside filtered air pushing through, any aerosol will be pushed away by air flow. Any possible remaining amplicon can be degraded by UV light or cleaned chemically and cabinet will be free from any amplicon. The question is what evidence you have or you can refer to show that usual lab hygiene is not sufficient to prevent PCR contamination. An amplicon that supposedly sticks to lab wall, how likely can be airborne again and land in a PCR tube. In a very slim probability if it lands how that really is going to affect results because the target amplicons will be in billions.

I strongly, would suggest external expert review of ancient DNA set up and practices if they can not control PCR contamination.


I never said PCR contamination is inevitable - there are many systems in place both molecular - (e.g using Uracil and UNG) and physical (ART tips and airflows) that minimise its potential to contaminate.

My warning was, and still is, around amplifying amplified DNA.

Using your scenario - working inside a PCR cabinet - as an example. If you open first round PCR products (to put thin into second round PCR) inside the cabinet aerosols are still an issue within the cabinet.

As some have pointed out; contamination is dependent on copy number - but in cases where 1st round PCR's have failed to amplify then any contaminating molecules can prime the 2nd round PCR. It is a risk, and people should be aware of it.

Evidence of contamination in PCR is everywhere - have you never had a contaminated PCR? - if the answer is yes then that is evidence that "control measures" you used failed. Thanks for the offer to review our lab practices - If you want to come visit our clean lab then please do;(https://jdlc.curtin.edu.au/facilities/trace.cfm) - always looking for cleaner ways to generate data.
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Old 11-22-2014, 02:53 AM   #18
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Thanks for clarifying your position on PCR contamination issues. I do not think that a mechanism or evidence has been provided for PCR contamination you have implied in earlier posts:
Quote:
Aerosols build in labs over time. In a post PCR area you can use ART tips, UV and gloves but this minimises contamination it won't remove it. A good PCR once 'opened' will generate aerosols with 10^5-10^9 copies of DNA that can travel through HEPA filters and build in a lab.
I was looking for evidence that shows an amplicon travels through HEPA filter, is not degraded by UV exposure, builds up in the lab and somehow ends up in another PCR reaction.
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Old 11-22-2014, 04:28 PM   #19
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Quote:
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Thanks for clarifying your position on PCR contamination issues. I do not think that a mechanism or evidence has been provided for PCR contamination you have implied in earlier posts:

I was looking for evidence that shows an amplicon travels through HEPA filter, is not degraded by UV exposure, builds up in the lab and somehow ends up in another PCR reaction.

Morning BioKiwi,

Sure, If you would like a few articles to help out with your research (it is not comprehensive - but provides a few exemplars).

I’d start with this paper;

http://www.plosone.org/article/info%....pone.0013042;

who comment;
"Carry-over contamination with products of previous PCR and cloning steps is one of the most serious threats for the generation of reliable results from minute quantities of DNA and also prevents the reliable evaluation of other contamination sources. The amplification and cloning of even a very small number of initial molecules produces up to 1013 molecules that are all identical and indistinguishable from those targeted. These contaminants may be carried over from previous amplification reactions due to aerosolization when the cap of a microtube is opened, and subsequent contamination of gloves, pipetting devices, laboratory surfaces, door knobs, handles of refrigerators and freezers, etc., in addition to reagents. This problem is exacerbated when semi-nested and nested PCR protocols are used. Carry-over contamination can be limited using dedicated devices, physical separation of the different experimental steps and stringent experimental procedures [51]. Used alone, these methods cannot guarantee complete protection [44], even when used in contained laboratories. Indeed, DNA is mostly spread by the experimenters who can be repeatedly contaminated by previous PCR and cloning products. These products can remain on many surfaces for long periods if they are not systematically identified and decontaminated after each potential contact with PCR products.”

and

"There is not a single decontamination method valid for all possible contamination sources occurring in PCR reagents and in the molecular biology laboratory and most common decontamination methods are not efficient enough to decontaminate short DNA fragments of low concentration”

UV clearly works to modify DNA but - even in hoods generates a low of shadows - it is not a ‘complete’ solution; see above and;

Comparison of the effects of sterilisation techniques on subsequent DNA profiling. http://www.ncbi.nlm.nih.gov/pubmed/17318649


HEPA - the physics of these filters are;

“DNA segments less than 300bp measure under 0.1 microns and pass through HEPA filters (0.3 micron pore size) in biological safety cabinets and are not eliminated by UV light or many disinfectant solutions.” - PCR for Clinical Microbiology: An Australian and International Perspective.


maybe finish with this paper on building clean labs and organising workflows.;
http://www.sciencedirect.com/science...40960211000732


PCR may be, to borrow your terminology “the corner stone of molecular biology” but within NGS workflows it is a necessary evil; bias, contamination, chimeras, error. You can go about designing workflows to minimise its impact - but " control" is an altogether different claim especially when amplifying amplified DNA.

If you want some more feedback on PCR contamination within NGS workflows maybe start a new thread on that topic? - I have aired my thoughts on the risks - the reality is that a lot of data on contamination does not end up in publications.
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